IDENTIFICATION and CHARACTERIZATION of Gatase1-LIKE Arac-FAMILY TRANSCRIPTIONAL REGULATORS in BURKHOLDERIA THAILANDENSIS

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2018 IDENTIFICATION AND CHARACTERIZATION OF GATase1-LIKE AraC-FAMILY TRANSCRIPTIONAL REGULATORS IN BURKHOLDERIA THAILANDENSIS. Adam Michael Nock University of Vermont

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IDENTIFICATION AND CHARACTERIZATION OF GATASE1-LIKE ARAC- FAMILY TRANSCRIPTIONAL REGULATORS IN BURKHOLDERIA THAILANDENSIS.

A Dissertation Presented

Adam Michael Nock

The Faculty of the Graduate College

The University of Vermont

In Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy Specializing in Microbiology and Molecular Genetics

May, 2018

Defense Date: March 14, 2018 Dissertation Examination Committee:

Matthew J. Wargo, Ph.D., Advisor John W. Barlow, D.V.M., Ph.D., Chairperson Jason W. Botten, Ph.D. Mary J. Dunlop, Ph.D. Keith P. Mintz, Ph.D. Cynthia J. Forehand, Ph.D., Dean of the Graduate College

ABSTRACT

The ability of bacteria to detect their surroundings and enact an appropriate response is critical for survival. Translation of external signals into a coherent response requires specific control over the transcription of DNA into RNA. Much of the regulation at this step is accomplished by transcriptional regulators, proteins that bind to DNA and alter gene expression. A wide-spread variety of regulators in bacteria is the AraC-family. These regulators are divided into two conserved domains and respond to a variety of compounds owing to different N-terminal domains. A subfamily of these regulators, GATase1-like AraC-family transcriptional regulators (GATRs), is described. These proteins contain an N-terminal domain with structural characteristics similar to enzymes that synthesize amine-containing compounds. Members of this subfamily of transcriptional regulators are found in a wide range of bacteria, however, few are characterized. A relatively high number of GATRs are encoded in the Burkholderia thailandensis genome. Therefore, we utilized this bacterium as a model to explore the function and diversity of these regulators.

GATRs in B. thailandensis divided into two groups based on bioinformatics analysis. The first group includes three members which we identified that contribute to the positive regulation of glycine betaine (GB) catabolism. GB can be utilized as a nutrient source or as a potent osmoprotectant. The regulation of this pathway in B. thailandensis differs from previously established models due to the interplay of these regulators. Homologs of two other GATRs in this group were identified that regulate carnitine and arginine catabolism. The second group of GATRs contains uncharacterized members with no known functions. A genetic strategy for engineering constitutive GATRs was developed and employed to investigate the transcriptional regulons of these GATRs. This approach yielded the identification of a novel GATR that represses expression of an operon producing a formaldehyde detoxification system, and is the first example of a GATR that functions as a repressor.

CITATIONS

Material from this dissertation has been published in the following form:

Nock A.M., Wargo M.J.. (2016). Choline Catabolism in Burkholderia thailandensis Is Regulated by Multiple Glutamine Amidotransferase 1-Containing AraC Family Transcriptional Regulators. Journal of Bacteriology, 198(18): 2503-14.

ACKNOWLEDGEMENTS

Any success that I have encountered, I must credit to the good fortune of a string of invaluable mentors. Although too long a list to properly enumerate, two in particularly must be mentioned here. Sohail Malik was instrumental in my training and essentially everything I know about biochemistry can be attributed directly to his instruction. His tutelage instilled both an appreciation for precise science and an orderly workspace.

Sohail’s attitude concerning science is as much curiosity as seriousness, and well- reflected the overall atmosphere at Rockefeller University, which is to continue the search for understanding the world around us. And without the guidance Thomas

Montville at Rutgers, I would likely not have made it to this crossroads at all. Tom’s lab solidified my interest in microbes while simultaneously helping me right my academic career and build a strategy to achieve future goals.

My parents James and Julia sacrificed a great deal of their comfort and interests in order to secure a better education and experience for John, Sarah, and myself. My wife

Audra has provided love and support throughout this venture. Her parents, Ben and

Grace, as well as our extended family, have been a continuous font of encouragement.

And in this time, somehow my dearest friends have not forgotten me no matter the distance. The generosity of the people in my life has been humbling.

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TABLE OF CONTENTS

Page ACKNOWLEDGEMENTS ...... iii

LIST OF FIGURES ...... ix

LIST OF TABLES ...... xi

CHAPTER 1: INTRODUCTION ...... 1

1.1 Sensing and responding to the environment ...... 1

1.2 Glycine betaine as a metabolite and signal ...... 2

1.3 The AraC family of transcriptional regulators ...... 3

1.4 Glutamine amidotransferase 1 (GATase 1)-containing AraC family

transcriptional regulators (GATRs) ...... 6

1.5 Burkholderia thailandensis as a model organism ...... 9

1.6 Focus of study and approach...... 14

1.7 Chapter 1 Figures ...... 16

1.8 Chapter 1 References ...... 28

CHAPTER 2: CHOLINE CATABOLISM IN BURKHOLDERIA THAILANDENSIS IS REGULATED BY MULTIPLE GLUTAMINE AMIDOTRANSFERASE1- CONTAINING ARAC-FAMILY TRANSCRIPTIONAL REGULATIORS ...... 36

2.1.1 Abstract ...... 37

2.1.2 Importance ...... 38

2.2 Introduction ...... 38

2.3 Methods...... 40 2.3.1 Culture conditions ...... 40

2.3.2 Strain Construction ...... 40 2.3.3 Growth assays ...... 41 2.3.4 Alignments and phylogenetic tree construction ...... 42 2.3.5 RNA-Seq ...... 42 2.3.6 qRT-PCR to confirm RNA-Seq findings ...... 44 2.3.7 Purification of GbdR1, GbdR2, and SouR...... 44 2.3.8 Electrophoretic mobility shift assays (EMSA) ...... 46 2.3.9 Generation of reporter constructs and β-galactosidase assays ...... 46

2.4 Results ...... 47 2.4.1 Organization of predicted B. thailandensis orthologs of P. aeruginosa choline catabolic genes ...... 47 2.4.2 gbdR1 and gbdR2 contribute differentially to choline catabolism ...... 49 2.4.3 GbdR1 and GbdR2 regulate transcription of genes involved in the choline catabolic pathway ...... 50 2.4.4 The glyA promoter is induced by choline under the control of GbdR1 and GbdR2 ...... 52 2.4.5 GbdR1 and GbdR2 bind directly to the glyA promoter ...... 54 2.4.6 B. thailandensis SouR regulates sarcosine metabolism ...... 55

2.5 Discussion ...... 57

2.6 Acknowledgements ...... 62

2.7 Chapter 2 Figures ...... 63

2.8 Chapter 2 Tables ...... 68

2.9 Chapter 2 References ...... 87

CHAPTER 3: EXPLORATION OF GATRS IN BURKHOLDERIA THAILANDENSIS ...... 91

3.1 Abstract ...... 91

3.2 Introduction ...... 92

3.3 Results ...... 95 3.3.1 Phylogenetic comparison of GATRs in Burkholderia ...... 95 3.3.2 Screening GATR functionality by carbon and nitrogen utilization ...... 97 3.3.3 Deletion of II1850 results in a catabolic defect on carnitine as a sole carbon Source...... 98 3.3.4 Deletion of I1774 results in a catabolic defect on arginine as a carbon source ...... 99 3.3.5 GbdR E36G constitutively upregulates transcription of members of the GbdR regulon...... 100 3.3.6 I0685 regulates expression of a formaldehyde dehydrogenase ...... 101 3.4 Discussion ...... 102 3.4.1 GATRs ...... 102 3.4.2 Identification of a GATR acting as a repressor of formaldehyde Detoxification ...... 104 3.4.3 II1850 is an ortholog of CdhR and regulates carnitine catabolism ..... 106 3.4.4 I1774 is an ortholog of ArgR and regulates arginine catabolism ...... 107 3.4.5 II1760, a potential regulator of cyanide detoxification ...... 108 3.4.6 Significance of choline-O-sulfate ...... 110 3.4.7 The absence of SouR in B. mallei ...... 110

3.5 Methods...... 110 3.5.1 Alignment of GATRs and phylogenetic tree construction ...... 111 3.5.2 Culture conditions for strains ...... 111 3.5.3 Construction of deletion strains ...... 111 3.5.4 Carbon and nitrogen utilization screens ...... 112 3.5.5 Prediction of amino acid mutations resulting in constitutively active GATRs ...... 113 vi

3.5.6 Generation of complement and constitutive GATR strains ...... 114

3.5.7 Growth of strains and RNA extractions ...... 114 3.5.8 Microarray analysis of constitutive mutants ...... 116 3.5.9 cDNA synthesis and quantitative real-time PCR ...... 116 3.5.10 Growth of I0685 deletion and complemented strains in the presence of Formaldehyde ...... 117

3.6 Chapter 3 Figures ...... 118

3.7 Chapter 3 Tables ...... 122

3.8 Chapter 3 References ...... 142

CHAPTER 4: DISCUSSION AND FUTURE DIRECTIONS ...... 148

4.1 A brief summary of progress ...... 148

4.2 Remaining questions about the regulation of glycine betaine catabolism in B.Thailandensis...... 149 4.2.1 What are the possibilities for GbdR1/GbdR2 interaction? ...... 149 4.2.2 Likely influences on the regulation of GB catabolism in B. thailandensis that remain unaddressed ...... 151 4.2.3 How important is GB as an osmolyte and are there pools of related molecules in B. thailandensis? ...... 152 4.2.4 How closely is the transport and catabolism of choline-O-sulfate linked to GbdR1 and GbdR2? ...... 154 4.2.5 The absence of SouR in B. mallei and potential effects on GB catabolism ...... 155 4.3 Deciphering the mechanism of GATRs ...... 157 4.3.1 How do GATRs interact with RNA polymerase? ...... 157 4.3.2 A novel mechanism for ATR function ...... 158 4.3.3 The potential functions of unknown GATRs ...... 159

4.4 Concluding remarks ...... 160 vii

4.5 Chapter 4 Figures ...... 162

4.6 Chapter 4 References ...... 164

COMPREHENSIVE REFERENCES ...... 169

APPENDIX A: BIOFILM FORMATION IN BURKHOLDERIA

THAILANDENSIS ...... 167

A.1 Introduction ...... 167

A.2 Methods ...... 168

A.3 Results and Discussion...... 171

A.4 Appendix A Figures ...... 176

A.5 Appendix A Tables ...... 177

A.6 Appendix A References ...... 182

APPENDIX B: A PROPHAGE AFFECTING B. THAILANDENSIS ...... 184

B.1 Introduction ...... 184

B.2 Methods ...... 185

B.3 Results and Discussion ...... 186

B.4 Appendix B Figure ...... 187

B.5 Appendix B Table ...... 189

B.6 Appendix B References ...... 192

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LIST OF FIGURES

Figure Page

Chapter 1 Figures Figure 1.1 - Structure of ATRs ...... 19 Figure 1.2 - The light-switch mechanism of AraC ...... 20 Figure 1.3 - Mechanism of RegA ...... 21 Figure 1.4 - Structure of GATRs ...... 22 Figure 1.5 - Distribution of GATRs ...... 23 Figure 1.6 - GATRs in B. thailandensis ...... 24 Figure 1.7 - Phylogeny of Burkholderia ...... 25 Figure 1.8 - Global endemicity of melioidosis ...... 26 Figure 1.9 - Beware of Burkholderia ...... 27

Chapter 2 Figures Figure 2.1- Comparison of B. thailandensis and P. aeruginosa choline metabolism and associated regulators...... 69 Figure 2.2 - Deletion of gbdR1 and gbdR2 result in altered growth on choline and its metabolites as sole carbon sources ...... 70 Figure 2.3 - The growth phenotypes of ΔgbdR1 and ΔgbdR2 can be complimented ..... 71 Figure 2.4 - The glyA promoter is induced by choline and its metabolites under the combined control of gbdR1 and gbdR2 ...... 72 Figure 2.5 - MBP-GbdR1 and MBP-GbdR2 bind to the regulatory region of the major GB catabolic operon (BTH_II1868-BTH_II1858) ...... 73 Figure 2.6 - SouR is a critical regulator of the choline catabolic pathway ...... 74 Figure 2.S1 - Complementation of ΔgbdR1ΔgbdR2 with gbdR1 or gbdR2 ...... 75 Figure 2.S2 - Validation of RNA-Seq experiments ...... 76

Figure 2.S3 - pAN27 is still responsive to choline in ΔsouR ...... 77 Figure 2.S4 - Usage of choline and sarcosine as sole carbon or nitrogen sources ...... 78

Chapter 3 Figures Figure 3.1 - GATR phylogeny based on full-length ...... 124 Figure 3.2 - GATR phylogeny based on GATase1-like domain ...... 125 Figure 3.3 - Growth of deletion strains on sole carbon sources ...... 126 Figure 3.4 - Growth of deletion strains on sole nitrogen sources ...... 127 Figure 3.5 - II1850 (cdhR) regulates carnitine catabolism ...... 128 Figure 3.6 - I1774 (argR) regulates arginine catabolism ...... 129 Figure 3.7 - Constitutive mutations in P. aeruginosa SouR overlaid on 3GRA ...... 130 Figure 3.8 - Constitutive GbdR rescues growth on choline ...... 131 Figure 3.9 - I0685 represses resistance to formaldehyde ...... 132 Figure 3.10 - II1912 synteny in other Burkholderia ...... 133 Figure 3.11 - FtrA is a potential regulator of cyanide detoxification ...... 134

Chapter 4 Figure Figure 4.1 - GATRs of known function in B. thailandensis ...... 163

Appendix A Figures A.1 - Searching for a link between phenotype and genotype ...... 178 A.2 - Biofilm phenotype subculturing in B. thailandensis E264 ...... 179

Appendix B Figure B.1 - A bacteriophage infecting B. thailandensis E264 ...... 190

LIST OF TABLES

TABLE Page

Chapter 2 Tables Table 2.1 - Transcript changes of B. thailandensis in response to choline ...... 79 Table 2.S1 - Strains and plasmids ...... 82 Table 2.S2 - Primers used in study ...... 83

Chapter 3 Tables Table 3.1 - Strain list ...... 135 Table 3.2 - Plasmid list ...... 136 Table 3.3 - Primer list ...... 137 Table 3.4 - Burkholderia GATR homologs based on gene synteny ...... 138 Table 3.5 - Predicted constitutive mutants ...... 139 Table 3.6 - GbdR E36G microarray results ...... 140 Table 3.7 - I0685 N32G microarray results ...... 141

Appendix A Tables Table A.1 - Strain Utilized in Study ...... 180 Table A.2 - RNAseq Analysis Correlation ...... 181

Appendix B Table Table B.2 - Sequencing results for cloned inserts...... 191

CHAPTER 1:

INTRODUCTION

1.1 Sensing and responding to the environment.

Microbes must be able to survey their molecular surroundings and respond accordingly in order to survive and compete in a changing environment. A desiccating or nutrient poor environment may prompt a bacterium to undergo a metabolic slowdown to conserve resources, or even build a resilient spore (1). Perhaps a bacterium has been fortunate enough to colonize a favorable environment and metaphorically stakes its claim by building a robust biofilm (2). But how exactly is this achieved? Some signals are detected through the membrane by two-component regulatory systems that consist of a histidine kinase and a response regulator, which transmit information through phosphorylation relays (3). Other molecules may diffuse or be actively transported into the cytosol, where they impact the physiology of the cell directly or be detected by one- component regulatory systems more commonly referred to as transcriptional regulators

(4). The response to these signals often relies on altering transcription or gene expression in the bacterium.

Transcription, the conversion of DNA into RNA by protein machinery, is a key to the central dogma of biology. DNA provides a master set of plans for the cell and a protected library of molecular information. The reading of the DNA and the production of RNA transcripts provide a translation of this information into functional directives for the cell to follow thus enabling it to persist and multiply. Control over transcription is a near universal point of regulation for the activities of all cells. A large portion of this

1 regulation is focused on the activities of the RNA polymerase holoenzyme, which is the protein machinery specifically responsible for the mechanistic process of transcription.

The RNA polymerase must be recruited to the DNA at specific sites to produce functionally useful RNA. Transcription regulators bind to DNA elements to specifically recruit to or obscure RNA polymerase binding sites. There are a wide range of transcriptional regulators, which are classified by the architecture and function of the constituent protein domains.

1.2 Glycine betaine as a metabolite and signal.

Glycine betaine (GB) is found in abundance in the natural environment. Part of the reason for this abundance of availability is the widespread utilization of precursor molecules that can be catabolized into GB. Choline, for example, is an important moiety for both phosphatidylcholine and sphingomyelin. These phospholipids are major components of the plasma membrane, particularly in some eukaryotes (5). Another precursor molecule for GB is carnitine. Carnitine is used by eukaryotes in the process of transporting lipids across the mitochondrial membrane (6). For these two reasons, GB is thought to be particularly abundant in the environmental context of eukaryotes. GB is also significant because it is utilized as an osmoprotectant, a compatible solute that protects the cell against salt stress, by many prokaryotes, eukaryotes, and archaea (7-11).

Many organisms can also metabolize GB for growth, using it as either a source of carbon or nitrogen (11). The abundance of GB and its metabolites in the environment may function as a signal to which bacteria can respond, especially in the context of association

2 with eukaryotes (11).

Pseudomonas aeruginosa is both able to utilize GB as an osmoprotectant, and contains a catabolic pathway that allows the bacterium to use GB as a nutrient source

(11). GB and related metabolites in P. aeruginosa also stimulate production of the extracellular phospholipase PlcH, a prominent virulence factor (12). GbdR (glycine betaine and dimethylglycine response regulator) is responsible for regulating the genes that catabolize choline into GB to dimethylglycine, and genes encoding proteins involved in the transport of these compounds (13). Two other closely related regulators also impinge on the regulation of catabolism related to GB in P. aeruginosa. SouR regulates the catabolic genes responsible for converting dimethylglycine to sarcosine and subsequently to glycine and serine (14). Additionally, the regulator for the carnitine catabolic and transport genes is CdhR (15). Carnitine provides an alternative metabolic route to GB. These three regulators share homology and are suggested to bind to the same or overlapping sequences in the promoter (14, 16). GbdR, SouR, and CdhR all belong to the AraC/XylS family of transcriptional regulators.

1.3 The AraC family of transcriptional regulators.

The AraC/XylS-family of transcriptional regulators (ATRs) is one of the most common types of prokaryotic transcriptional regulators. The namesake of this family,

AraC, is responsible for the regulation of the araBAD operon that contains genes for the catabolism of arabinose (17-19). ATRs are defined by the homologous ninety-nine amino acid residues in the C-terminus of the ATR (20). The C-terminus is characterized by a helix-turn-helix DNA-binding domain, followed by a linker region joined to the N-

3 terminal region of the protein, typically containing ligand binding and dimerization capabilities (21-23) (Figure 1.1). ATRs regulate multiple or single operons, and are often themselves encoded divergently from genes they regulate. ATRs target genes that encode catabolic enzymes, virulence factors, secondary metabolism, transport, and others (24).

AraC itself exists as a dimer at normal concentrations, whether or not it is bound to DNA and or arabinose (25). AraC regulates the L-arabinose operon by the ‘light switch mechanism’ (Figure 1.2). The operon is activated when each monomer binds an arabinose molecule, which increases affinity for the two proximal half sites, I1 and I2, activating transcription. In the absence of arabinose, one monomer binds I1 while the other binds to an alternate half site (O2) which is further upstream, resulting in repression of transcription by DNA looping (26). The I1 and I2 half site sequences are composed of direct repeat symmetry, as opposed to other transcription factors which have inverted repeats (27). This well-studied example has contributed to our general understanding of

ATRs. However, not all of these insights are broadly applicable to the rest of this family of regulators.

The other canonical example of ATRs is XylS, which regulates alkyl benzoate catabolism and was discovered on the TOL plasmid of Pseudomonas putida (28). Unlike

AraC, which makes some contacts with the σ70 subunit of RNA polymerase, XylS may utilize either σ70 or σ54 at its promoters (28, 29). Another difference is that while it also binds to similarly structured half sites and activates as a dimer, XylS typically exists in solution as a monomer (30). XylS and other dimerizing ATRs (e.g. RegA) only bind target promoters in the presence of the appropriate ligand rather than utilizing the ‘light- switch mechanism’ (31) (Figure 1.3). Still other ATRs like MarA, SoxS, and Rob, which

4 are involved in E. coli stress response and drug resistance, are examples of ATRs that bind promoters in respective regulons as monomers (32).

Many of the differences in ATRs stem from differences in the N-terminal domain of the protein that imparts ligand specificity and dimerization capacity. Notably, MarA and SoxS lack the ligand binding domain characteristic of many other ATRs altogether

(33). The N-terminal domains of ATRs are derived from the binding domains of other protein families. An ATR in Sinorhizobium meliloti contains a cupin-like domain that enables a direct interaction with cyclic-di-GMP, quite different from binding sugars as a ligand (34). Still others utilize ligands such as urea and bicarbonate (35, 36).

The expression of genes coding for ATRs is often tightly regulated in order to exert fine-tuned control over the respective regulons. As regulators, the number of molecules per cell is often not significantly more than the number of target promoters, as is the case for AraC (27, 37). SoxS occurs at approximately 100 molecules per cell while impacting the regulation of about one hundred genes (38). XylS concentration per cell has also been experimentally determined and is in the range of 200 molecules per cell, while its target genes may vary in number due to variable copy number of the TOL plasmid (30). Many ATRs, including AraC, participate in some variety of auto-regulatory scheme. MarA, an ATR involved in multidrug resistance in pathogenic Escherichia coli, has also been demonstrated to bind to its own promoter (39). The turnover rate of MarA is only several minutes, allowing the TetR-family repressor MarR, which is located in the same operon, to bind in the absence of MarA ligand (33). SoxS and Rob, are ATRs that regulate genes in overlapping pathways with MarA and compete for similar promoters thus impacting the regulation of MarA induced genes including MarA itself (32).

The transcriptional control of ATR regulons is often also modulated by other types of transcriptional regulators, either directly or indirectly. The cyclic-AMP receptor protein (CRP), which is involved in the regulation of many genes in E. coli, is required for AraC-mediated induction of the L-arabinose catabolic operon (40). CRP is suggested to contribute to the relief of the repressive DNA looping caused by AraC as well as assist in the recruitment of RNA polymerase through direct contacts (27). ATRs are subjected to anti-activation from a family of proteins designated AraC negative regulators (ANRs)

(41). These proteins interact with ATRs directly to prevent DNA binding. ANRs also interact with HN-S family proteins, which are responsible for DNA condensation and the silencing of AT-rich sequences (42). HN-S family proteins may also directly impact promoter regions of ATR targets, or the promoters of the ATRs themselves (43).

1.4 Glutamine amidotransferase 1 (GATase 1)-containing AraC family transcriptional regulators (GATRs).

We identified a subset of ATRs with common homology that we refer to as

GATRs. GATRs contain two C-terminal helix-turn-helix motifs like other ATRs, but the

N-terminal domain bears homology to Class I GATases (Figure 1.4), a family of enzymes that catalyzes the transfer of amide groups to various substrates. The distribution of these regulators is quite broad, including both Gram-positive and Gram-negative bacteria

(Figure 1.5), however there are few characterized representatives of this type of ATR.

These GATRs are particularly common amongst the Proteobacteria. The genomes of many members of the Proteobacteria contain multiple GATRs, rather than just one or two. Interestingly, GATRs are also found in eukaryotes including the pharaoh ant, pea

6 aphid, and Entamoeba histolytica (44). Many GATRs are labeled as FtrA or FtrA-like, owing to a described GATR in Caulobacter crescentus, which is suggested to be a putative regulator under limited oxygen conditions of an alternative branch of the thiamine synthesis pathway, myo-inositol degradation, and two aminotransferases (45).

Analysis of amino acid sequence homology and synteny of adjacent genes reveals

GATRs designated FtrA are unlikely to be orthologous and likely to have differential functions. Some GATRs contain a conserved cysteine residue within the N-terminal domain, which is a characteristic of the GAT superfamily of which GATases are part.

Members of the GAT superfamily of proteins are typified by a variable strand number Rossmann fold (46), which is series of parallel β-sheets suggested to play a role in nucleotide binding (47). This superfamily includes a wide variety of proteins with well characterized functions aside from the GATases. These include proteases like Pfp1 from the Archean Pyrococcus furiosus (48), and the chaperone Hsp31 in E. coli (49). The yeast homolog of Hsp31 in addition to chaperone activity, has a methylglyoxal detoxification activity under conditions of oxidative stress (50). Many members of the GAT superfamily appear to be evolutionary derivatives of ThiJ, a bacterial protein involved in the synthesis of thiamine (51). This includes the human protein DJ-1, which is protective against oxidative stress in the brain and associated with Parkinson’s disease (52, 53). The

GAT superfamily also shares evolutionary features with catalases, and typical sub- families may differ by the size of a variable region between strands of the Rossmann fold and occasional lack of the typical catalytic triad, cysteine-histidine-aspartate/glutamate

(46). The superfamily is rounded out by particular members of the AraC-family (GATRs) and the GATases.

GATases are known to be involved in the biosynthesis of a wide variety of aminated molecules including amino acids, aminated sugars, nucleotides, and more (54).

GATases typically utilize glutamine as the nitrogen donor, but some utilize primarily exogenous ammonia instead of or in addition to glutamine. The only noted exception is

Glu-tRNAGln amidotransferase, which also utilizes asparagine at a low rate (55).

GATases are divided both functionally and topologically into two domains of a monomer or as heterodimers of translationally separate subunits (56, 57).

The first domain, referred to as the GAT (glutamine amidotransferase) domain, consists of both the amine binding and amindotransferase activity. The second domain contains the synthase activity and is generally quite divergent owing to the chemically diverse amide receptors participating in the reaction to generate a particular product. The liberated ammonia is shuttled through a molecular channel between the amidotransferase and synthase domains (58-60).

The GAT domain is divided into two distinct classes, as defined by sequence homology (61). Class 1 has a conserved catalytic triad consisting of a cysteine, histidine, and glutamine. Class 2 members possess a single highly conserved cysteine in the amino terminus. The remaining amino acid residues of the catalytic triad are less well conserved.

The first characterized GATR was ArgR in P. aeruginosa. ArgR regulates the arginine succinyltransferase pathway of arginine catabolism (62). Although P. aeruginosa has three additional pathways for arginine utilization, it is unable to use arginine as a sole carbon source in the absence of argR (63). In P. aeruginosa, ArgR also plays a critical role in the catabolism of lysine. This is due to the requirement of ArgR for

8 expression of the ldcA gene which codes for a decarboxylase critical for the catabolism of lysine and its use as a sole carbon source (64). ArgR is unrelated to the similarly named, hexameric regulation protein found in E. coli and other enterics despite regulating similar components of the metabolic pathway (65, 66). The remaining recently characterized

GATRs are implicated in different branches of GB catabolism in P. aeruginosa.

1.5 Burkholderia thailandensis as a model organism.

GATRs are especially prevalent in Burkholderia. B. thailandensis in particular possesses thirteen bioinformatically predicted GATRs (Figure 1.7). Because many examples of GATR-containing microorganisms only have one or two putative GATRs, a sample size of thirteen represents a significant survey of GATRs with respect to their different functions. For this reason, and the fact that B. thailandensis is a genetically tractable model organism for which there are preexisting genetic tools (67-69), we have concentrated our efforts on exploring this subset of GATRs in order to achieve a greater understanding of GATRs as a family.

Burkholderia thailandensis is a saprophytic, β-proteobacteria associated with the rhizosphere. The genome is arranged as two circular chromosomes possessing 5712 genes over 6.7 megabases (strain E264) (70). This genome is larger compared to familiar bacteria such as E. coli K-12 at 4.6 megabases, which indicates an increased number of genes based on the relatively linear correlation between genome size and the number of genes in bacterial species (71). A larger number of genes suggest an enhanced ability to take advantage of a larger diversity of compounds found in the environment, and is typical of free-living bacteria isolated from the environment (72). Initially, Burkholderia

9 species fell under the genus Pseudomonas due in part to metabolizing a similar array of compounds and filling a similar environmental niche. Burkholderia are now recognized as an independent genus (73), and have undergone several reclassifications in the scientific literature (Figure 1.6). More recently, a large number of primarily environmental Burkholderia have been reclassified and placed in the new genus,

Paraburkholderia (74).

Burkholderia thailandensis was originally isolated from rice paddy soil samples in Thailand and recognized as distinct from the pathogenic Burkholderia pseudomallei due to its ability to catabolize arabinose and an apparent lack of clinical cases caused by any isolates with this ability (75-77). It is considered avirulent, and classified as a biosafety level 1 organism by many research institutions. Burkholderia species also tend to be naturally resistant to many antibiotics due to their contingent and variety of antimicrobial pumps and capsules (78). The metabolic flexibility of many

Burkholderiales has made them a focus for studies in bioremediation (79). Burkholderia species are also notable for their capacity to produce secondary metabolites, including antifungal compounds (80, 81).

Utilizing B. thailandensis as a model organism has the advantage of serving as a substitute for its more pathogenic relatives due to its close genetic similarity (70). The

Burkholderia genus can be roughly subdivided into two further distinct clades. The first group includes Burkholderia pseudomallei and Burkholderia mallei, potent pathogens and select agents. It is to this group that B. thailandensis is most closely related and the three organisms are thought to share a core metabolic genome (70). In addition, B. thailandensis also possesses many genes associated with virulence including fimbriae,

10 several capsule operons, several type III secretion systems, several type-VI secretion systems, and flagella suited to both extracellular and intracellular lifestyles (82). The presence of these virulence-associated genes in B. thailandensis may be related to the antagonistic and symbiotic interactions of these and other Burkholderiales have with different amoeba species (83-85). Members of this clade are also able to stimulate the fusion of different phagocytic cells into multinucleated giant cells in both humans and arthropods (86-88). The second group includes Burkholderiales that are often members of the rhizosphere or plant pathogens. This second group is also notable for including the namesake organism of the genus, Burkholderia cepacia, a pathogen of the onion plant that infiltrates the onion causing a tannish to yellow ooze of bacteria and rendering it inedible (89) (Figure 1.8A). However, these organisms may cause opportunistic infections in humans, particularly in immune compromised individuals. The best example of this opportunistic pathogenicity is in patients with cystic fibrosis (CF) where many of these Burkholderiales make up what is known as the Burkholderia cepacia complex

(Bcc) and greatly increase morbidity (90). Similarly to the Pseudomallei group, B. thailandensis shares a large portion of its genes with members of the Bcc, but has the advantage of genetic tractability as the genomes of Bcc members are notoriously genetically malleable due to recombination between insertion elements, and are consequently less facile for genetic experimentation (91).

As a model for B. pseudomallei is particularly advantageous because B. thailandensis is significantly less virulent in mammals, and therefore less hazardous for laboratory work (92, 93). Although B. thailandensis retains the ability to live intracellularly in eukaryotic cell culture, the lack of a major capsule representing a

11 primary virulence factor in B. pseudomallei, allows the bacterium to be rapidly cleared by the innate immune system (94). Ultimately, the only incidents of disease attributed to B. thailandensis are a car accident were an infant was ejected from the vehicle into a drainage ditch (95), a traumatic leg amputation due to a motorcycle incident (96), and an unusual case of fever in an elderly man in Chongqing, China (97). In all of these cases, infection caused by B. thailandensis does not seem to be the primary health concern and the ability to identify the causative agent in a clinical setting can be limited by available resources and ability to isolate the correct bacterium.

B. pseudomallei is the causative agent of melioidosis, a disease that is rapidly gaining recognition in developing countries. Melioidosis, also known as Whitmore’s disease after it’s discoverer Alfred Whitmore (98) (Figure 1.8B), is difficult to diagnose in part because it has similar symptoms to other common illnesses and many countries where it is endemic lack the clinical facilities needed to accurately identify it (99). The infection has often progressed to an advanced stage by the time it is identified, often when abscesses are detectable on the spleen and other internal organs when the patient already may be septic. In addition, B. pseudomallei infection may be latent for an indeterminate amount of time before becoming active. The exact means by which this occurs is unknown, but may have to do with the ability of B. pseudomallei to infiltrate immune cells and survive intracellularly. Due to this ability, the illness was previously known as the “Vietnam Time Bomb”, as American veterans of the Vietnam War might not become ill until years after returning home with one documented case occurring after sixty years. This phenomenon was particularly prevalent among helicopter crews, perhaps due to the generation of dust particles upon takeoff and landing (100). There are

12 a predicted 165000 cases of melioidosis per year with an approximately fifty percent mortality rate based on computer models (101). Infection may manifest a number of symptoms and result in abscesses and septic shock (102). Cases of melioidosis correspond to seasonal monsoons, possibly due to the resulting rising water table or the disturbance of the soil (103, 104). B. thailandensis and B. pseudomallei may directly compete for the same environmental niche, as isolation of these organisms in

Northeastern compared to other regions of Thailand favors B. thailandensis and correspond with a lower level of melioidosis cases (105) (Figure 1.9). Additionally, B. pseudomallei and B. thailandensis both can compete experimentally with their multiple contact dependent inhibition systems, type-IV secretion systems, in addition to other systems (106-108).

The select agent status of B. pseudomallei and B. mallei also contribute to the difficulty of working with these pathogens. B. mallei is an extremely close relative of B. pseudomallei that has been genetically reduced putatively due to the high presence of transposons in its genome. As a consequence, B. mallei is an obligate intracellular pathogen and has never been isolated from soil. It is the causative agent of glanders in equines (horses, donkeys, mules) (Figure 1.8C), although it can also infect humans who have been in contact with infected equines (109). Due to efforts to eradicate infected animals, the last natural case of glanders in the United States occurred in 1937. However, it is still endemic in other countries where the use of equines as beasts of burden is common (109). B. mallei was actively used as an agent of biological warfare during

World War I by Germany in an effort to sabotage the supply of horses to Europe from the

United States (110). B. pseudomallei was also researched biological weapon by the

13 infamous Japanese unit 731 during World War II were the notable ability of B. pseudomallei to persist in pure water for long periods of time was exploited to contaminate well water in China (111). The potential of B. pseudomallei for use as a biological weapon was also researched by the Soviet Union as part of their notorious

Biopreparat biological weapons program (112). Due to this colorful history, the politics and legal restrictions on studying these pathogens impose great monetary costs and hurdles to research. This makes the avirulent B. thailandensis an attractive, cost-effective alternative model organism.

1.6 Focus of study and approach.

The work in this thesis focuses on understanding the role of GATRs in the context of B. thailandensis. Because of the importance of GB catabolism, we first sought to identify an ortholog for GbdR (113). In Chapter 2 we identify multiple orthologs of

GbdR and describe an alternative model for regulation of the GB catabolic pathway.

Regulons for the identified GATRs are determined through RNA-Seq, reporter assays, and electro mobility shift assays. Chapter 3 describes our efforts to identify the function of additional GATRs in B. thailandensis. Strains with constitutive GATRs were generated by combining structural data with information about known constitutive mutations in a characterized GATR. This allowed for the determination of the regulon of an unknown GATR, I0685. I0685 is the first GATR found to act as a repressor, and prevents expression of a formaldehyde detoxification system. Two other GATRs were characterized by screening B. thailandensis GATR deletion strains against a small library

14 of compounds used in growth assays as either a sole carbon or sole nitrogen source.

Finally, Chapter 4 explores the significance of these findings, what unanswered questions remain, and suggests future experiments to expand on these studies.

1.7 Chapter 1 Figures

Figure 1.1 – General Representation of ATRs.

AraC-family transcriptional regulators are separated into an N-terminal and C-terminal domain, joined by a short linker. The N-terminal domain includes both effector binding and dimerization capacity. A short N-terminal arm, which affects the nature of the dimerization interaction, is also present. The C-terminal domain imparts DNA binding ability and consists of two helix-turn-helix (HTH) motifs.

Figure 1.2 – The light-switch mechanism of AraC.

Figure adapted from Yang et al. (31). (A) The schematic of AraC and the ‘light-switch mechanism’ are a model for the understanding of ATRs. (B) The AraC dimer binds to the upstream element O2 and I1 of the araBAD promoter in the absence of arabinose ligand. (C) In the presence of ligand, the N-terminal arm changes arrangement, affecting the dimerization interaction and preference for half sites to the I1 and I2 sites, making the complex capable of activating transcription.

Figure 1.3 – Mechanism of RegA.

Figure adapted from Yang et al. (31). (A) In the absence of ligand, RegA remains

- unbound. (B) In the presence of its ligand, HCO3 (bicarbonate), RegA binds to and activates its target promoters.

Figure 1.4 – General Representation of GATRs.

GATRs are markedly similar to other ATRs, consisting of similar functional domains.

However, the N-terminal domain is homologous to glutamine amidotransferase class 1 enzymes (GATase1).

Figure 1.5 – Distribution of GATRs.

(A) Protein architecture used for Conserved Domain Architecture Retrieval Tool

(CDART) query to find GATRs in the NCBI non-redundant protein sequence database

(44). (B) Result of query showing the broad presence of GATRs in bacteria, particularly proteobacteria, and several examples in eukaryotes.

Figure 1.6 – Phylogeny of Burkholderia.

The Burkholderia genus may be broken up into a group containing pathogens similar to

B. pseudomallei, a group encompassing opportunistic pathogens that include members of the BCC, and phytopathogens. Many classical members of the genus can now be separated into the new, but closely related genus, Paraburkholderia. This phylogenetic tree was generated using PATRIC (114).

Figure 1.7 – GATRs in B. thailandensis.

GATRs, and rough schematics of their genomic context bioinformatically, predicted in B. thailandensis by BLAST homology to GbdR from P. aeruginosa. GATRs are depicted in magenta, while other predicted genes are depicted in yellow.

Figure 1.8 – Beware of Burkholderia.

Members of the Burkholderia genus may cause a variety of diseases in plants and animals. (A) Example of sour skin of onion caused by B. cepacia (115). (B) Medical examples of cutaneous melioidosis in patients (116). (C) A horse afflicted with Glanders, caused by B. mallei. Source: The Brooke Hospitals for Animals

(https://www.thebrooke.org/our-work/india/glanders-impact-horse-owners)

Figure 1.9 – Global endemicity of melioidosis.

Figure adapted from Center for Disease Control website. Cases of melioidosis are generally concentrated near the tropical regions, particularly in South East Asia.

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CHAPTER 2:

Choline catabolism in Burkholderia thailandensis is regulated by multiple glutamine

amidotransferase1-containing AraC-family transcriptional regulators

Adam M. Nock1 and Matthew J. Wargo1,2

1Department of Microbiology and Molecular Genetics and 2The Vermont Lung Center,

University of Vermont College of Medicine, Burlington, VT 05405

2.1.1 Abstract

Burkholderia thailandensis is a soil-dwelling bacterium that shares many metabolic pathways with the ecologically similar, but evolutionarily distant,

Pseudomonas aeruginosa. Among the diverse nutrients it can utilize is choline, metabolizable to the osmoprotectant glycine betaine and subseqeuntly catabolized as a source of carbon and nitrogen, similar to P. aeruginosa. Orthologs of genes in the choline catabolic pathway in these two bacteria showed distinct differences in gene arrangement as well as an additional orthologous transcriptional regulator in B. thailandensis. In this study, we showed that multiple glutamine amidotransferase1 (GATase1)-containing

AraC-family transcription regulators (GATRs) are involved in regulation of the B. thailandensis choline catabolic pathway (gbdR1, gbdR2, souR). Using genetic analyses and sequencing the transcriptome in the presence and absence of choline, we identified the likely regulons of gbdR1 (BTH_II1869) and gbdR2 (BTH_II0968). We also identified a functional ortholog for P. aeruginosa souR, a GATR that regulates the metabolism of sarcosine to glycine. GbdR1 is absolutely required for expression of the choline catabolic locus, similar to P. aeruginosa GbdR, while GbdR2 is important to increase expression of the catabolic locus. Additionally, the B. thailandensis SouR ortholog (BTH_II0994) is required for catabolism of choline and its metabolites as carbon sources, whereas in P. aeruginosa, SouR function can by bypassed by GbdR. The strategy employed by B. thailandensis represents a distinct regulatory solution to control choline catabolism and thus provides both an evolutionary counterpoint and an experimental system to compare the acquisition and regulation of this pathway during environmental growth and infection.

2.1.2 Importance

Many proteobacteria that occupy similar environmental niches have horizontally acquired orthologous genes for metabolism of compounds useful in their shared environment. The arrangement and differential regulation of these components can help us understand both the evolution of these systems and the potential roles these pathways have in the biology of each bacterium. Here we describe the transcriptome response of

Burkholderia thailandensis to the eukaryote-enriched molecule choline, identify the regulatory pathway governing choline catabolism, and compare the pathway to that previously described for Pseudomonas aeruginosa. These data support a multi-tiered regulatory network in B. thailandensis, with conserved orthologs in the select agents

Burkholderia pseudomallei and Burkholderia mallei, as well as the opportunistic lung pathogens in the Burkholderia cepacia clade.

2.2 Introduction

Burkholderia thailandensis is a saprophytic, soil dwelling bacterium common in tropical and subtropical regions, and is an opportunistic pathogen of insects, plants, nematodes, and amoeba (1-3). B. thailandensis is used as a less virulent model for the select agents B. pseudomallei and B. mallei, the causative agents of melioidosis and glanders, respectively. The reduced virulence of B. thailandensis is primarily due to the absence of the major capsule locus important for B. pseudomallei virulence in mammals

(4), although it is used to study the effect of type III section systems on phagocytic escape and retains other genes associated with virulence (5, 6). Despite virulence

38 differences, a great deal of the core genome is shared between B. thailandensis and its relatives, including many pathways for accessory metabolism.

Genes predicted to be involved in choline catabolism are found throughout the B. pseudomallei group (BPG) and Burkholderia cepacia complex (BCC) (7, 8). Many soil bacteria can use choline as a sole carbon and nitrogen source, and this catabolic pathway may be particularly important for bacteria associated with eukaryotes (9). Choline is part of the polar head group of both phosphatidylcholine and sphingomyelin, which together constitute the majority of lipids on the outer leaflet of eukaryotic cell membranes (10), and are also abundant in pulmonary surfactant (11). In addition to its role as a nutrient source, choline metabolism can generate glycine betaine (GB), an important osmoprotectant (12) and inducer of virulence factor production (13). The conversion of choline to GB has been shown to be important for Escherichia coli survival in urine (14) and P. aeruginosa survival in the mammalian lung (15, 16). Many organisms, including

Burkholderiales, maintain an intracellular GB pool, potentially as a hedge against future osmotic stress (17). Choline can also be found in the rhizosphere, exuded from plant roots, likely to influence populations of bacteria in this environment (18). In fact, many plants accumulate high levels of GB as an osmoprotectant, notably beets from whence the appellation ‘betaine’ was derived (19). In the environment, secreted or decaying organic matter provides a metabolic opportunity that holds much promise to microbial opportunists poised to exploit them.

Our lab has previously studied the choline catabolic pathway and its regulation in

Pseudomonas aeruginosa (20-22), and examination of orthologous genes in B. thailandensis suggested an alternate, perhaps more complex, regulatory strategy. In P.

39 aeruginosa, the critical regulator of GB catabolism is GbdR, a type-I glutamine amidotransferase (GATase1)-like AraC-family transcription regulator (GATR) that regulates the genes required for sequential demethylation of GB to glycine in P. aeruginosa (Fig 2.1A, left side) (22). In contrast to P. aeruginosa, B. thailandensis and other Burkholderiales appeared to have two GbdR orthologs (Fig 2.1B, C), whose co- occurrence and conserved synteny suggested that they could play non-overlapping roles in B. thailandensis choline catabolism. In this study, we have investigated the contributions of the two B. thailandensis GbdR orthologs to choline and GB catabolism and to global gene expression in response to choline. These data and follow-up analyses reveal that the choline catabolism pathway in B. thailandensis, while bearing many similarities to P. aeruginosa, contains key differences in organization and regulation.

2.3 Methods

2.3.1 Culture conditions.

Burkholderia thailandensis E264 (ATCC® 700388™) (23) cultures were inoculated from -80 oC glycerol stocks onto LB (10 g/L tryptone, 5 g/L yeast extract, 5 g/L NaCl) with 1.5% agar and incubated at 37 oC overnight. For protein expression and molecular cloning, Escherichia coli DH5α or E. coli T7 express cells (New England

BioLabs (NEB)) were then grown in LB broth containing 100 µg/mL carbenicillin, kanamycin, or trimethoprim (Tp), as appropriate.

2.3.2 Strain Construction.

Deletion strains (Table S1) were generated as described in Thongdee et al. and the resistance markers were removed using the methods described in Choi et al. (24, 25).

Briefly, a PCR product was amplified via splice overlap extension PCR that consisted of a trimethoprim resistance marker amplified from pUC18mini-TN7T-Tp, including the flanking Flp recombinase sites, and with regions of homology upstream and downstream of the gene of interest (See primers in Table S2). The resultant PCR product was then used to naturally transform B. thailandensis that had been grown in defined media (DM) as previously described [24]. To remove the resistance marker, pFLPe2 was electroporated into the Tp resistant strain and plated onto low-salt LB (LSLB: 10 g/L tryptone, 5 g/L yeast extract, 3 g/L NaCl, 1.5% agarose, pH 7.5 with 250 µg/mL zeocin and 0.2% L-rhamnose for Flp recombinase expression). Cells were cured of pFLPe2 by growth at 42 oC. Resultant colonies were streaked onto plates with zeocin, Tp, or without antibiotics to identify colonies where the trimethoprim marker and pFLPe2 had been lost.

The ΔgbdR1, ΔgbdR2, and ΔsouR phenotypes were complemented by cloning each gene and ~250 bps of the upstream regulatory region into pUC18mini-TN7T-Zeo and integration into one of the two attTn7 sites in B. thailandensis (26, 27) (Table 2.S1).

Appropriate control strains were generated by attTn7 integration with the empty pUC18mini-TN7T-Zeo vector (Table 2.S1).

2.3.3 Growth assays.

Growth assays were performed by starting overnight cultures in MOPS minimal media supplemented with 20 mM pyruvate and 5 mM glucose incubated at 37 oC (28).

Cultures were diluted in MOPS media to an OD600 of 0.07 and 30 µL were used to

41 inoculate 470 µL of MOPS media supplemented with carbon sources as described in a flat bottom 48-well plate (Costar). These plates were incubated at 37 oC with agitation and OD600 measured using a Biotek Synergy 2 plate reader.

2.3.4 Alignments and phylogenetic tree construction.

Sequence alignments and phylogenetic analysis were preformed using MEGA version 6 (29). Amino acid sequences of the relevant GATRs were aligned using

MUSCLE (30). The ftrA sequence, a GATR homolog from Escherichia coli, was included as an outgroup to root the phylogenetic tree.

2.3.5 RNA-Seq.

For RNA-Seq, wild type, ΔgbdR1, ΔgbdR2, and ΔgbdR1Δgbdr2 strains were streaked from frozen stocks onto LB agar plates and incubated overnight at 37 oC. These plates were used to inoculate 3 mL MOPS with 20 mM pyruvate and 5 mM glucose cultures which were incubated for 18 hours at 37 oC on a rotary wheel. To initiate the experiment, 1 mL of these cultures was added to 2 mL of pre-warmed MOPS with 30 mM pyruvate or 2 mL MOPS with 30 mM pyruvate and 1.5 mM choline; resulting in final concentrations of 20 mM pyruvate and 1 mM choline. These cultures were incubated for four hours at 37 oC. Cells from 1.5 mL of culture were collected by centrifugation and the resultant pellets were resuspended in RNA Protect Bacteria

Reagent (Qiagen) and incubated at room temperature for 10 minutes. Cells were collected by centrifugation and resuspended in 50 µL 10 mM Tris, 1 mM EDTA (TE) buffer with 3 mg/mL lysozyme and incubated at room temperature for 5 minutes. To each of these

42 resuspensions, 1 µL 20 mg/mL proteinase K, 0.5 µL of 10% SDS, 1 µL of DNase I

(RNase-free, Ambion), and 2 µL of 50 mM MgCl2 were added and samples were incubated for 5 minutes at room temperature. RNA was then prepared using the Qiagen

RNeasy kit according to the manufacturer’s instructions. Eluate from this purification was further treated with 11 µL 10x DNase I buffer and 2 µL DNase I (RNase-free,

Ambion) and incubated at 37 oC for 20 minutes to remove contaminating DNA and re- purified using the Qiagen RNeasy protocol with the optional on-column DNase I step.

The resulting purified RNA was assessed for purity by PCR and quantified by bioanalyzer chip, depleted of 16S and 23S rRNA by using MICROBExpress™ Bacterial mRNA Enrichment (Ambion), and subsequently reassessed by Bioanalyzer (Agilent

Technologies). RNA-Seq DNA libraries were prepared by the Vermont Cancer Center –

College of Medicine Massively Parallel Sequencing Facility using an Illumina TruSeq

Stranded Total RNA Library Prep Kit and were run on an Illumina Hi-Seq 1500 to generate single end reads. Read quality was checked with FastQC (31). Samples were aligned to the B. thailandensis E264 chromosomes I (NC_007651) and II (NC_007650) using BWA (32), duplicate reads were marked using Picard (version 1.110; Broad

Institute of MIT and Harvard, [http://broadinstitute.github.io/picard/]) , and gene counts were calculated using summarizeOverlaps from the GenomicRanges package (33) before differential expression was called using the DESeq package (34) in R (version 3.0.1; R

Development Core Team, [https://www.r-project.org/]) with assistance from the

University of Vermont College of Medicine Bioinformatics Shared Resource group.

RNA-Seq data has been deposited in NCBI GEO under record GSE81652.

2.3.6 qRT-PCR to confirm RNA-Seq findings.

Wild-type, ΔgbdR1, ΔgbdR2, and ΔgbdR1Δgbdr2 strains were grown and induced as described for the RNA-Seq experiment. RNA was extracted similarly to the

RNA-Seq, except that a third DNase I treatment was performed using the Ambion DNA- free kit (AM1906) as per the manufacturer’s protocol. cDNA was generated using

Superscript IV First-Strand Synthesis System (Invitrogen) using 24 ng of RNA combined with the 5’-NSNSNSNSNS-3’ primer previously described (35). qPCR was performed as described in Willsey et al. (36). Briefly, 0.2x SYBR Green I nucleic acid gel stain

(Thermo Fisher Scientific (TFS)) was used with Q5 High-Fidelity 2x master mix (NEB) and amplified using cycle conditions of 98 oC for two minutes, 98 oC for 20 seconds, annealing at 60 oC for 30 seconds, and extension at 72 oC for 20 seconds repeated by going back to the second step 39 times. Reactions were performed in technical duplicate and biological triplicate with RT primers designed for BTH_I1140 (rplU), BTH_II1869

(gbdR1), BTH_II1861 (gbcA), BTH_II1853 (putative porin), BTH_II0968 (gbdR2),

BTH_II0966 (cbcV), BTH_II0994 (souR), and BTH_II0996 (soxB). A dilution series was used to generate a standard curve for each primer set. Each reaction was normalized to its respective rplU value, and then induced values were divided by uninduced values to derive a fold effect value.

2.3.7 Purification of GbdR1, GbdR2, and SouR.

gbdR1, gbdR2, and souR were each cloned into the pMAL-C2X expression vector generating an N-terminal maltose binding protein tag. The constructs were transformed

44 into E. coli T7 Express, transformants selected on LB agar + 100 µg/mL carbenicillin, and positive colonies used to inoculate 20 mL cultures of LB + 100 µg/mL carbenicillin.

After 18 hours, these cultures were used to inoculate 500 mL flasks of LB + 100 µg/mL

o LB carbenicillin, grown to an OD600 of 0.3, and induced for 4 hours at 30 C with 500

µM IPTG. Cells were collected by centrifugation. Cells were then washed, collected by centrifugation, and the resulting cell pellets were flash frozen in liquid nitrogen. Cell pellets were resuspended on ice in 10 mL of chilled chromatography and lysis buffer (20 mM Tris pH 7.4, 500 mM NaCl, 20 mM imidazole, 1x HALT protease inhibitor (TFS)).

10 µL DNase I was added to the resuspended cell pellet, and the mixture was incubated for 10 minutes on ice. The resuspended cells were lysed with a French cell press using three 1100 PSI passages and NP-40 was then added to a final concentration of 0.05%.

The lysate was clarified by centrifugation for 15 minutes at 15,000 rpm at 4 oC. The supernatant was passed over a column containing 1 mL bed volume of amylose resin beads (NEB). The column was then washed with 10 mL of column buffer 300 (20 mM

Tris pH 7.4, 300 mM KCl, 1 mM EDTA, 0.05% NP-40, 0.7 µL/mL β-mercaptoethanol), then 10 mL column buffer 150 (20 mM Tris pH 7.4, 150 mM KCl, 1 mM EDTA, 0.05%

NP-40, 0.7 µL/mL β-mercaptoethanol). Protein was eluted in column buffer 150 containing 10 mM maltose and twelve 300 µL fractions were collected. Fractions were analyzed by SDS-PAGE acrylamide gels stained with Coomassie. Since GATR proteins tend to precipitate at higher concentrations, fractions of lower concentration were pooled and dialyzed in Slide-A-Lyzer® Dialysis Cassette G2 (TFS) against 2 L column buffer

150. Aliquots were flash frozen in liquid nitrogen and stored at -80 oC.

2.3.8 Electrophoretic mobility shift assays (EMSA).

Biotinylated or unlabeled primers were used to generate DNA probes from appropriate templates containing the putative promoter regions using Q5® High Fidelity

DNA polymerase (NEB), then subjected to QIAquick PCR Purification Kit (Qiagen) and quantified using a Nanodrop 1000 (TFS). 20 µL reactions were assembled using 2x binding buffer (10 mM Tris pH 7.4, 200 mM KCl, 2 mM EDTA, 2 mM DTT, 10% glycerol, and 200 µg/µL bovine serum albumin (37)), 5 ng/µL poly (dI-dC), 25-75 nM purified MBP-tagged protein, 1 nM biotinylated DNA probe, and optionally 20 nM unlabeled competitor DNA probe. Reactions were incubated in a 30 oC waterbath for 20 minutes, then 5 µL 5x EMSA loading buffer (1x TBE, 20% glycerol, 0.01% xylene cyanol, 0.01% bromophenol blue) was added and samples were loaded onto a 5% acrylamide 0.5x TBE gel. Gels were run for 1 hour at 100 volts then transferred to a

BioDyne-B nylon membrane (Pierce) for 1 hour at 80 volts in 0.5x TBE at 4 oC. DNA was crosslinked to the membrane using a UV Stratalinker 2400 (Stratagene) on the auto crosslink setting, and the biotin-labeled probe visualized using the Pierce LightShift

Chemiluminescent EMSA Kit (TFS) and imaged on a ChemiDoc XRS+ molecular imager (Bio-Rad). Density of resulting bands was quantified using Quantity One software from Bio-Rad.

2.3.9 Generation of reporter constructs and β-galactosidase assays.

Yeast homologous recombination was used to replace the kanamycin resistance gene in the shuttle vector pMQ131 (38) (apaHA3) with the Tp resistance gene (dhrFII)

46 resulting in pAN1. Yeast recombination was also used to replace the lacZα fragment with the full lacZYA operon resulting in pAN7. The intergenic region between gbdR1 and glyA, as well as truncations of this region, were ligated into the SphI site proximal to the lacZYA ribosomal binding site, and transformants selected on LB agar + 100 µg/mL

Tp at 37 oC. Tp resistant colonies were used to inoculate 3 mL MOPS media with 20 mM pyruvate, 5 mM glucose, and 100 µg/mL Tp, which were incubated for 48 hours at 37 oC.

Cells from these cultures were collected by centrifugation at 13000 rpm for 2 min at room temperature, washed with 1 mL MOPS media, collected again by centrifugation and the pellets resuspended in MOPS media. 30 µL of this resuspension was used to inoculate each well in a 48-well plate that contained 470 µL MOPS media with 20 mM pyruvate and with or without 1 mM choline. These cultures were incubated for 48 hours at 37 oC on an orbital shaker at 170 rpm in a humidified chamber. The extended induction time was shown to be necessary for detectable β-galactosidase activity empirically in our hands, and as indicated by Kang et al. (39). β-galactosidase activity was quantified as previously described (21).

2.4 Results

2.4.1 Organization of predicted B. thailandensis orthologs of P. aeruginosa choline catabolic genes.

The choline catabolic pathway and its regulation in P. aeruginosa have been described previously (20, 40). Briefly, in P. aeruginosa the TetR family transcription factor BetI represses expression of betIBA and is derepressed by choline, allowing production of the BetA and BetB enzymes that to convert choline into GB (41-43). The

GATase1-like AraC-family transcription regulator (GATR) GbdR responds to GB and dimethylglycine (DMG) by inducing expression of catabolic genes including gbcA-B, the dgc operon (PA5376, PA5377, dgcA, dgcB, PA5400, PA5401), and the sarcosine oxidase genes, which together contribute to the sequential demethylation of GB to DMG, sarcosine, and finally glycine (Fig 2.1A,C). SouR, also a GATR, controls transcriptional induction of the sarcosine oxidase genes in response to sarcosine (36). Homologs of betI and betBA in B. thailandensis have been described previously (39).

Sequence analysis revealed two likely GbdR orthologs in B. thailandensis,

BTH_II1869 (gbdR1) and BTH_II0968 (gbdR2), with strong amino acid sequence homology to P. aeruginosa GbdR (48.2% and 47.0% identity and 75.5% and 71.3% similarity, respectively, using BLOSUM50 along the entire protein length). Alignments of the characterized P. aeruginosa GATRs (ArgR, CdhR, SouR), and their putative B. thailandensis orthologs, revealed that the B. thailandensis GbdR orthologs were more similar to each other than to P. aeruginosa GbdR or any other GATRs (Fig 2.1B). This suggested that they might also have similar functions or play separate roles within the same catabolic pathway. The presence of a more distantly related GATR divergently transcribed from the sarcosine oxidase operon (Fig 2.1B, C), led us to predict that

BTH_II0994 was a functional ortholog of P. aeruginosa SouR, though the phylogenetic tree suggests that they share no recent paralogy.

On the P. aeruginosa chromosome, gbdR is located one gene away from the cbcXWV transport genes (Fig 2.1C), previously implicated in the transport of choline and its immediate metabolites into the cytosol from the periplasm (44), and adjacent to the sdaB gene known to participate in the pathway (45). In B. thailandensis, gbdR1 is located

48 adjacent to a putative operon containing the bulk of predicted orthologous genes encoding enzymes needed for GB catabolism to sarcosine, while gbdR2 is present within a locus containing the putative cbcXWV orthologs (Fig 2.1C). The homology of GbdR1 and GbdR2 with P. aeruginosa GbdR, combined with their chromosomal locations adjacent to genes involved in choline metabolism, strongly suggested that both transcription factors were involved in the regulation of this pathway. The synteny of the gbdR1 and gbdR2 loci in B. thailandensis E264 is conserved throughout the various clades of Burkholderiales, including species such as B. pseudomallei, B. mallei, B. cenocepacia, B. ambifaria, and B. multivorans. This conservation in synteny suggests that both GbdR1 and GbdR2 play roles in the regulation of choline metabolism in

Burkholderia.

2.4.2 gbdR1 and gbdR2 contribute differentially to choline catabolism.

B. thailandensis, like P. aeruginosa (46), can use choline as a sole source of carbon and/or nitrogen (Fig 2.2A). Based on our predictions of gbdR1 and gbdR2 involvement in choline metabolism we tested the ability of each single deletion strain and the double deletion strain to grow using choline as a sole carbon source. Wild type B. thailandensis grows on choline as a sole carbon source, while a ΔgbdR1 or

ΔgbdR1Δgbdr2 strain cannot (Fig 2.2A, right side). Deletion of gbdR2 alone resulted in reduced growth on choline as a sole carbon source (Fig 2.2A, right side). On plates, none of the manipulations appeared to alter growth on pyruvate as a sole carbon source (Fig

2.2A, left side). The impaired but visible growth of the ΔgbdR2 strain led us to examine the growth kinetics of these strains on choline and its downstream metabolites GB and

DMG. Deletion of gbdR2 results in slower growth than wild type on choline, GB, and

DMG, comprised of both an extended lag phase and a lower growth rate during exponential phase (Fig 2.2B). Deletion of these transcription factors did not substantially alter growth kinetics using pyruvate as a sole carbon source, but did alter maximal yield in liquid culture in the double deletion strain (Fig 2.2B), although this difference was not apparent on solid media (Fig 2.2A, left side). The ΔgbdR1 and ΔgbdR1Δgbdr2 strains showed no measurable growth on any of the choline-related metabolites tested within 100 hours.

Integration of gbdR1 at the attTn7 site restored growth of a ΔgbdR1 strain on choline, GB, and DMG (Fig 2.3A). Growth in GB and DMG was very similar to wild type, while growth of the complement strain on choline exceeded that of wild type on choline (Fig 2.3A). Similarly, integration of gbdR2 at the attTn7 site restored growth of a

ΔgbdR2 strain on choline-related carbon sources and resulted in more rapid growth than in wild type for choline, GB, and DMG (Fig 2.3B). We also tested complementation of

ΔgbdR1ΔgbdR2 with either gbdR1 or gbdR2. The ΔgbdR1Δgbdr2 attTn7::gbdR1 strain was able to grow on choline as a sole carbon source in MOPS minimal media, though not at wild type levels, whereas ΔgbdR1Δgbdr2 attTn7::gbdR2 was not capable of growth on choline (Fig 2.S1). Taken together, these data suggest that gbdR1 and gbdR2 regulate the choline catabolic pathway, with gbdR1 absolutely required and gbdR2 playing an accessory role.

2.4.3 GbdR1 and GbdR2 regulate transcription of genes involved in the choline catabolic pathway.

To identify the GbdR1 and GbdR2 regulons, we exposed our deletion strains to minimal media with pyruvate as a primary carbon source and with or without 1 mM choline, and analyzed the transciptomes by RNA-seq. Exposure of wild-type B. thailandensis to choline resulted in 57 transcripts being induced or repressed more than

2.3 fold, many predicted to be involved in quaternary amine catabolism and transport

(Table 2.1). A prominent cluster of induced genes from BTH_II1868 to BTH_II1853 includes orthologs to the P. aeruginosa glyA (BTH_II1868), the dgc operon

(BTH_II1867, BTH_II1866, BTH_II1865, BTH_II1864, BTH_II1863, and BTH_II1862), gbcA (BTH_II1861), gbcB (BTH_II1860), and betX (BTH_II1859). Based on these predictions, the putative operon consisting of glyA to BTH_II1858 (Fig 2.1C) encodes the genes likely to be responsible for the breakdown of GB to sarcosine. Also induced was a nearby predicted operon consisting of BTH_II1855- BTH_II1853, encoding likely orthologs of P. aeruginosa betC, cosX, and a predicted porin. Another GATR,

BTH_II0994, and the divergently transcribed operon consisting of BTH_II0995 (sdaA-2),

BTH_II0996 (soxB), BTH_II0997 (soxD), BTH_II0998 (soxA), BTH_II0999 (soxG), and

BTH_II1000 were also induced and we hypothesized that these are the B. thailandensis souR and the sarcosine oxidase genes responsible for the breakdown of sarcosine to glycine (36).

Homologs of cbcXWV in P .aeruginosa, an operon important for the transport of choline, are present in B. thailandensis and are also induced by choline. The gbdR2 gene is situated in this operon, between BTH_II0967 (cbcW) and BTH_II0969 (cbcX). The aforementioned genes are induced by choline in the wild type strain, but are not in either

ΔgbdR1 or ΔgbdR1Δgbdr2, suggesting an absolute requirement of gbdR1 in the

51 regulation of these genes. Cells lacking gbdR2 express all of the genes in the major catabolic operon, glyA-BTH_II1858, in response to choline but to a lower level than the wild type. Interestingly, the relative induction of the transcripts in the transport operon containing gbdR2 are unaffected by the absence of gbdR2, suggesting that there is no autoregulation of the gbdR2 operon.

To confirm our RNA-Seq results, we preformed qRT-PCR on a subset of induced genes and the transcription factors of interest, normalized to rplU transcript levels (Fig

S2). The wild type B. thailandensis response to choline in our qRT-PCR experiment reflected our RNA-Seq results. gbcA, cbcV, and soxB are all induced in response to choline, suggesting overall induction of their putative operons. The qRT-PCR data also shows that the GATRs gbdR1 and souR transcripts were slightly induced in response to choline while gbdR2 was induced at a higher level. This slight increase in gbdR1 expression observed may be negligible and therefore it is unclear if induction of gbdR1 is a mechanism by which GbdR2 regulates choline catabolism. Overall, we interpret the matching trends in expression between the RNA-Seq and qRT-PCR experiments as a validation of our findings.

2.4.4 The glyA promoter is induced by choline under the control of GbdR1 and

GbdR2.

pAN27 is a reporter plasmid containing the full 408 bp intergenic region between gbdR1 and glyA, such that the glyA promoter drives lacZYA expression. This reporter was used to assess the transcriptional control of the glyA promoter. This putative promoter likely controls the expression of BTH_II1868 to BTH_II1857 and thus governs the bulk

52 of genes needed to convert GB to sarcosine. In a B. thailandensis wild-type background, pAN27 lacZ expression is induced by choline, GB, and DMG, but not ethylcholine (Fig

2.4A). Ethylcholine is a non-metabolizable inducer of GbdR-dependent transcription in

P. aeruginosa (40). The inability of this choline analog to elicit similar effects in a

GbdR/GbdR2 dependent promoter suggests specificity differences between GbdR and

GbdR1 or GbdR2, a metabolite transport difference, or a difference in the specificity of choline oxidase. No β-galactosidase activity was detected when pAN27 in B. thailandensis was induced using 1 mM sarcosine, similar to findings in P. aeruginosa for gbdR-dependent induction of choE, a choline esterase (40), or for the phospholipase plcH

(47, 48) (data not shown).

ΔgbdR1, ΔgbdR2, and ΔgbdR1Δgbdr2 deletion strains carrying pAN27 failed to show significant choline-induced β-galactosidase induction when compared to the wild type control (Fig 2.4B), and basal expression was also low. This suggests that both gbdR1 and gbdR2 are required for robust induction of the glyA promoter. We would have predicted that ΔgbdR2 would produce some β-galactosidase activity in response to choline, as ΔgbdR2 can use choline as a sole carbon source (Fig 2.2). The incongruity may be due to the unusually long time it takes to observe β-galactosidase activity in B. thailandensis, combined with the long delay in growth of ΔgbdR2 on choline (Fig 2.2).

As a follow-up experiment, we transformed pAN27 into a ΔsouR to determine if SouR is required for glyA promoter induction in B. thailandensis (Fig 2.S3). β-galactosidase activity in ΔsouR in response to choline is robust, higher than in wild type, potentially due to blockage of the catabolic pathway and subsequent buildup of intermediate metabolites. This indicates that SouR is not required for induction of glyA, but could also

53 point to positive feedback control between sarcosine metabolism and regulation of upstream steps.

Reporter plasmids derived from pAN27 with progressively smaller portions of the intergenic region between gbdR1 and glyA were produced (-340, -251, -147, -85 with respect to the glyA putative translational start site) in order to map the choline-dependent portion of the glyA regulatory region. Exposure of cells containing these constructs to choline resulted in choline-dependent induction of β-galactosidase activity for -340 and -

251, but not -147 or -85 (Fig 2.4C). This suggests that the region from -251 bp to the translation start site of glyA is necessary for a response to choline.

2.4.5 GbdR1 and GbdR2 bind directly to the glyA promoter.

N-terminally tagged MBP-GbdR1 and MBP-GbdR2 were produced and assayed by EMSA using biotinylated DNA probes representing sections of the intergenic region between gbdR1 and glyA. Probe 1 is 216 bp covering the -320 to -104 bp region with respect to the glyA translational start site, while probe 2 is 178 bp covering the -160 to

+18 region. MBP-GbdR1 could bind probe 1 in a concentration-dependent manner, and binding to biotinylated probe 1 could be significantly competed with unlabeled probe 1

(Fig 2.5A). MBP-GbdR1 did not interact with probe 2 (right-most lane, Fig 2.5A), and unlabeled probe 2 was markedly less able to compete with MBP-GbdR1 binding to labelled probe 1. These data support a GbdR1 binding site within probe 1. Similar results were obtained using MBP-GbdR2, which was also able to bind to probe 1, not probe 2, and could be similarly competed from labeled probe 1 using unlabeled probe 1 (Fig

2.5B), and again much less so with the unlabeled probe 2. The shift produced by MBP-

GbdR2 consists of multiple bands that are likely the result of oligomerization. Results using 6xHis-N-terminally tagged versions of GbdR1 and GbdR2 produced the same results and banding patterns, although the solubility of the 6-His version was much lower than the MBP fusions (data not shown). These results provide evidence that both GbdR1 and GbdR2 can both bind to the glyA promoter. When combined with results from our reporter assays (Fig 2.4), we infer that the binding site for GbdR1 and GbdR2 is likely to be contained within the -251 to -160 bp region with respect to the glyA translational start site. As we have previously shown for P. aeruginosa GbdR and SouR, addition of their presumptive ligand did not impact binding to DNA (data not shown).

2.4.6 B. thailandensis SouR regulates sarcosine metabolism.

Sarcosine is a downstream metabolite of choline and orthologs of P. aeruginosa sarcosine oxidase genes were identified in our RNA-Seq results (Table 2.1, Fig 2.1C). P. aeruginosa uses sarcosine as a sole carbon and nitrogen source and this is regulated by the transcription factor, SouR (36). While B. thailandensis can use choline as a sole carbon or sole carbon and nitrogen source, B. thailandensis is unable to grow on sarcosine as a sole carbon source, or as a sole carbon and nitrogen source (Fig 2.S4). We hypothesize that the ability of B. thailandensis to transport extracellular sarcosine may be limited in comparison with P. aeruginosa, and the rate of import is insufficient to support growth as a sole carbon source. It is, however, able to utilize sarcosine as a sole nitrogen source when pyruvate is available as a primary carbon source, and does so in a SouR dependent manner (Fig 2.6A). Deletion of gbdR1 or gbdR2 has no major effect on the ability of B. thailandensis to utilize sarcosine as a nitrogen source, but ΔsouR does show

55 a reduced level of growth.

BTH_II0994 is a GATR divergently transcribed from the sarcosine oxidase operon and we hypothesized that it might be functionally orthologous to souR in P. aeruginosa (Fig 2.1C), despite BTH_II0994 and SouR not sharing a high degree of similarity when compared to the other GATR orthologous pairs (Fig 2.1B). To test if

BTH_II0994 was a functional souR, we generated a B. thailandensis ΔsouR strain and used it to determine if souR is required for growth on choline. ΔsouR fails to grow on choline, GB, or DMG as a sole carbon source (Fig 6B). This is contrary to regulation in

P. aeruginosa, where choline, GB, and DMG can still be used as sole carbon sources for growth in the absence of souR, although at a diminished growth rate (36). This result also suggests that GbdR1 and GbdR2 are insufficient to cause induction of the sarcosine oxidase genes by themselves or together, contrary to the regulation scheme in P. aeruginosa (Fig 2.1A). This phenotype can be complemented by chromosomal integration of souR at an attTn7 site under the control of its native promoter (Fig 2.6B).

MBP-SouR was produced and its DNA binding assessed using EMSA with a biotinylated 230 bp oligo designated probe 3, covering the region -208 to +22 base pairs relative to the putative translation start site of sdaA-2, the first gene in the likely sarcosine oxidase operon of B. thailandensis. MBP-SouR was able to bind the biotinylated probe 3, but not the probe 2 negative control. The binding of MBP-SouR to Biotinylated probe 3 could be competed off with unlabeled probe 3, but not unlabeled probe 2, suggesting that this interaction is specific (Fig 2.6C). The binding of MBP-SouR to this putative promoter region coupled to the phenotype of the ΔsouR strain suggests that SouR is responsible for the regulation of the sarcosine oxidase genes in B. thailandensis.

2.5 Discussion

The ability to utilize GB as a compatible solute, providing protection against osmotic and other stresses without creating significant disruption of normal cell processes, is nearly ubiquitous among bacteria, however the ability to catabolize GB is not as widespread (49). B. thailandensis and other species in this genus possess the enzymes for utilizing choline and its metabolites as carbon and nitrogen sources, taking advantage of this widely available biomolecule as befits their description as metabolically adaptable bacteria. P. aeruginosa, another soil-dwelling and metabolically adaptable microorganism, shares this ability and was our original model for the study of the GB catabolic pathway and its regulation (20). B. thailandensis and P. aeruginosa inhabit similar environments, possess similarly sized genomes, encode diverse metabolic pathways, and are thus considered to have similar generalist strategies, despite B. thailandensis and P. aeruginosa belonging to the β-proteobacteria and ɣ-proteobacteria classes, respectively. The metabolic enzymes in the choline catabolic pathway are well conserved and the orthologous genes in different species are readily identifiable.

However, the identification of gbdR2 and the alternate gene organization in B. thailandensis prompted us to ask if the choline regulatory network was as well conserved.

We show here that B. thailandensis uses an alternate regulatory solution to control this pathway compared to P. aeruginosa. Both B. thailandensis and P. aeruginosa can store pools of GB (17) and metabolize it as a carbon and nitrogen source, therefore the different regulatory networks suggest evolution of alternate strategies to control the decision to store or catabolize.

Although the data presented here indicates that choline catabolism is regulated differently in B. thailandensis than in P. aeruginosa (Fig 1A), the basic components of the pathway are conserved and thus the overall scheme of the pathway is the same. In both organisms, choline catabolism begins with choline-dependent de-repression of the catabolic genes responsible for the conversion of choline to GB, betBA, mediated by the choline-sensing TetR-family transcription factor BetI. These genes have been described previously (41), and evidence suggests they function in B. thailandensis as they do in P

.aeruginosa (39). In P. aeruginosa GB is subsequently metabolized to DMG by GbcA and GbcB heterodimer (50, 51), and from DMG to sarcosine by enzymes in the dgc operon (52), all under the regulation of GbdR (20). In B. thailandensis the same enzymatic steps are under the control of GbdR1, modulated by GbdR2. Sarcosine is then demethylated to glycine by components of the sarcosine oxidase operon and glyA1, controlled in tandem by GbdR and SouR in P. aeruginosa, and controlled separately by

SouR, and GbdR1 and GbdR2 respectively in B. thailandensis as described below.

In this study we determined that both GbdR1 and GbdR2 participate in the regulation of GB catabolism, and characterized their regulons. Both regulators are

GATRs with strong amino acid sequence similarity to GbdR in P. aeruginosa, and deletion strains confirmed that both play roles in the regulation of choline catabolism and catabolism of its downstream metabolites (Fig 2, 3). Combining the transcriptional data with the growth phenotypes of the deletion strains supports gbdR1 as required for expression of the major catabolic cluster of genes and the transport operon that includes gbdR2. gbdR2, on the other hand, is important for a robust response to choline, enhancing expression of many genes required to import and catabolize choline and its metabolites,

58 particularly the major catabolic cluster (Fig 2.1C, Table 2.1). The operon containing gbdR2 and the putative cbcWVX orthologs, is likely regulated solely by GbdR1. Although the presence of gbdR1 is sufficient for growth (Fig 2.2A-B), reporter assays and growth studies showed that both gbdR1 and gbdR2 are required for a robust transcriptional response (Fig 2.4A) and growth on choline (Fig 2.2). In agreement with promoter mapping results (Fig 2.4C), EMSA data suggests that GbdR1 and GbdR2 likely share a similar binding site, raising the question of whether or not they compete for the same binding site or act synergistically. A possibility is that GbdR1 and GbdR2 possess differential DNA binding Kd values that result in differential response to choline and its metabolites under different physiological conditions.

In addition to gbdR1 and gbdR2, our transcriptome analysis revealed a third

GATR induced in response to choline, BTH_II0994, which we confirmed as a functional souR ortholog in B. thailandensis. In P. aeruginosa, SouR regulates the expression of the sarcosine oxidase catabolic operon, the components of which are responsible for converting sarcosine to glycine (36). SouR in B. thailandensis is less similar to GbdR,

GbdR1, and GbdR2 than is the SouR in P. aeruginosa (Fig 1B). B. thailandensis souR

(BTH_II0994) is divergently transcribed from the sarcosine oxidase operon, whereas P. aeruginosa souR (PA4184) is distantly located and part of a two gene operon. These factors suggest a divergence in how souR was acquired or evolved in both organisms.

The inability of B. thailandensis to grow on sarcosine as a sole carbon source was surprising given that choline is readily catabolized and the enzymes for sarcosine catabolism are expressed (Fig 2.6A). We found that souR in B. thailandensis is required for sarcosine catabolism, unlike in P. aeruginosa where GbdR can induce the sarcosine

59 oxidase operon in the absence of souR (36). The complementation and EMSA for the sarcosine oxidase operon putative promoter provides further evidence that BTH_II0994 is indeed the functional souR ortholog in Burkholderia (Fig 2.6B, C). It is not clear if SouR is capable of autoregulation of souR or if GbdR1 controls its expression directly.

GbdR1, GbdR2, and SouR are all GATRs implicated in the regulation of genes involved in the catabolism of quaternary amines or their metabolites. GATRs are members of the AraC transcription factor family, grouped by their canonical C-terminal

DNA binding domain, but unlike classic AraC proteins they contain a GATase1-like N- terminal domain. The GATase1-like domain is characterized by its homology to class-I glutamine amidotransferases, which bind glutamine or ammonia and participate in the amidation/deamidation reaction (53, 54). Many of the GATR N-termini, notably excluding SouR in P. aeruginosa, also retain a bioinformatically identified cysteine residue that would be part of the functional catalytic triad of GATase1 family enzymes.

The N-terminal domain of AraC proteins is typically involved in ligand binding and dimerization, which is a characteristic important for their functionality, and dimerization is often affected by the binding of ligand (55). P. aeruginosa possesses other GATRs implicated in the catabolism of N-substituted amines including argR, the regulator of arginine catabolism (56), and cdhR, the regulator of carnitine catabolism (20). Putative argR and cdhR orthologs are also present in B. thailandensis (unpublished data). We hypothesize that the GATase1-like domain has been combined with the AraC-style DNA binding domain to enable detection of charged amine-containing compounds and that

GATRs as a whole may represent a sub-family of transcription factors that regulate metabolism of accessory nitrogen sources, including choline and its metabolites.

The transport of choline and derivative molecules also appears to be handled differentially between B. thailandensis and P. aeruginosa based on genomic information.

P. aeruginosa possesses an array of BCCT and ABC-family transporters that have been implicated in choline and GB transport that are effective under different osmotic conditions or primarily utilized when choline is in such abundance that it can be used as a carbon or nitrogen source (44). B. thailandensis possesses fewer transporters, with only one putative homolog of the BCCT-family transporter (BTH_II1109), compared to the three in P. aeruginosa (betT1, betT2, betT3). B. thailandensis possess a putative amino acid permease, BTH_II1858, as part of the major GB catabolic operon with no obvious ortholog in P. aeruginosa. The placement of BTH_II1858 and conserved synteny among

Burkholderia suggests a role as a transporter for choline or its derivatives, but there is no functional evidence as of yet. Some Pseudomonads have a putative ortholog of

BTH_II1858 identified as an ethanolamine transporter, ethanolamine being similar to choline in its role as a head group for phospholipids, suggesting the possibility of a link to general fatty acid metabolism. Both B. thailandensis and P. aeruginosa also possess orthologs of the choline/GB transporter, opuC that was described in P. syringae and found to function under hyperosmolar conditions (57). B. thailandensis orthologs to many of the P. aeruginosa periplasmic binding proteins that mediate ABC-transporter dependent import of quaternary amines (cbcX (BTH_II0969), caiX (BTH_II1849), betX

(BTH_II1859), cosX (BTH_II1853)), but these are apparently not sufficient for efficient sarcosine transport (Fig 2.S4). This result is perhaps not surprising as limited current data suggest that sarcosine does not compete well with choline for these transporters in P. aeruginosa (58). To date the sarcosine transporter has not been identified, and the

61 functional differences between P. aeruginosa and B. thailandensis for sarcosine utilization in the background of a similar metabolic pathway may provide a platform for identification of this transporter.

In this study we have identified an alternative model for the regulation of choline catabolism that incorporates multiple GATRs and examined their respective regulons.

The conservation of gbdR1 and gbdR2 throughout the Burkholderia genus suggests that differences in regulation of choline catabolism outlined here represent a model that can be extrapolated to the more pathogenic strains, as well those strains associated with the rhizosphere. Contrasting the models of choline catabolism in B. thailandensis and P. aeruginosa will serve as a useful tool to probe the remaining unanswered questions concerning the pathway.

2.6 Acknowledgements

We would like to thank Jeff Bond, Julie Dragon, Robert Devins, and David J

Shirley for bioinformatics assistance. We also owe thanks to Graham Willsey and Lauren

Hinkel for critical reading of the manuscript.

2.7 Chapter 2 Figures

Figure 2.1 - Comparison of B. thailandensis and P. aeruginosa choline metabolism and associated regulators.

(A) Model of choline catabolic regulation in P. aeruginosa (left) and B. thailandensis

(right). (B) Phylogenetic tree of relevant GATRs in B. thailandensis and P. aeruginosa rooted to FtrA, a GATR present in E. coli. (C) Diagram of the chromosomal context of gbdR, gbdR1, gbdR2, souR and major catabolic genes for the catabolism of GB and subsequent metabolites for B. thailandensis and P. aeruginosa.

Figure 2.2 - Deletion of gbdR1 and gbdR2 result in altered growth on choline and its metabolites as sole carbon sources.

(A) Growth of B. thailandensis deletions strains on MOPS minimal media agar plates supplemented with either 20 mM pyruvate or 20 mM choline and incubated at 37 oC for

72 hours. Δgbdr1 and ΔgbdR1ΔgbdR2 fail to grow on choline while ΔgbdR2 exhibits less growth. (B) Growth curves reveal the inability of Δgbdr1 and ΔgbdR1ΔgbdR2 to grow on choline or its metabolites while ΔgbdR2 exhibits a slow-growth phenotype on choline, glycine betaine, and dimethylglycine. Growth curves are representative of at least three experiments per condition at each time point, each with biological triplicates, and the error bars represent standard deviation.

Figure 2.3 - The growth phenotypes of ΔgbdR1 and ΔgbdR2 can be complimented.

(A) Complementation of ΔgbdR1 at attTn7 with gbdR1 and its putative promoter results in restoration of ability to grow on choline and its metabolites. (B) Complementation of

ΔgbdR2 at attTn7 with gbdR2 results in an increased growth rate on choline and its metabolites, surpassing the growth rate of wild type. Growth curves are representative of at least three experiments per condition at each time point, each with biological triplicates, and the error bars represent standard deviation.

Figure 2.4 - The glyA promoter is induced by choline and its metabolites under the combined control of gbdR1 and gbdR2.

(A) β-galactosidase activity from pAN27 is induced in the presence of choline and its metabolites, but not pyruvate. (B) gbdR1 and gbdR2 are both required for significant β- galactosidase induction in the presence of choline. (C) Promoter truncations of the glyA promoter reveal that the critical region is located between -147 and -340 with respect to the predicted translational start site of glyA. In each case, data presented represents the standard error of the mean determined from three independent experiments, each with three technical replicates. Statistical notation in panel (A) based on one-way ANOVA with Dunnett’s post-test using the pyruvate group as the comparitor, while (B) & (C) were analyzed by two-way ANOVA with Sidak’s post-test comparing with vs. without choline. Abbreviations and symbols: pyruvate (Pyr), choline (Cho), glycine betaine (GB), dimethylglycine (DMG), ethylcholine (EtCho), not significant (n.s.), p < 0.05 (*), p <

0.01 (**), p < 0.001 (***), p < 0.0001 (****).

Figure 2.5 - MBP-GbdR1 and MBP-GbdR2 bind to the regulatory region of the major

GB catabolic operon (BTH_II1868-BTH_II1858).

(A) MBP-GbdR1 binds to biotin-probe 1 in a dose dependent manner (lanes 3-5). The interaction is specific as it does not bind to biotin-probe 2 (lane 8) and can be competed off biotin-probe 1 with unlabeled probe 1 (lane 6 compared to lane 4, 17.2% shift versus

75.2% shift respectively), but less so with unlabeled probe 2 (lane 7 compared to lane 4,

51.5% shift versus 75.2% shift respectively). (B) MBP-GbdR2 also binds specifically to probe 1 in a dose dependent manner (lanes 3-5) while failing to bind to biotin-probe 2

(lane 8). The binding between MBP-GbdR2 and biotin-probe 1 is also specific, as it can be competed by unlabeled probe 1 (lane 6 compared to lane 4, 4.3% shift versus 16% respectively) but less so by unlabeled probe 2 (lane 7 compared to lane 4, 7.7% shift versus 16% respectively).

Figure 2.6 - SouR is a critical regulator of the choline catabolic pathway.

(A) B. thailandensis can utilize choline as a nitrogen source. Strains were grown on 20 mM Pyruvate with either 5 mM or choline (C) or sarcosine (S) for 72 hours at 37 oC.

ΔgbdR1, ΔsouR and ΔgbdR1ΔgbdR2ΔsouR were significantly diminished in their ability to utilize choline as a nitrogen source. ΔgbdR2 utilized both choline and sarcosine as a nitrogen source at similar levels to wild type. ΔsouR used sarcosine as a nitrogen source at reduced rates compared to the wild type. OD600 measurements were normalized to basal growth without a nitrogen source and then normalized to wild type and represented as percent growth. (B) Deletion of souR prevents B. thailandensis from using choline as a carbon source. Complementation of souR at an attTn7 site with souR, under the control of

65 its putative native promoter, restores the ability of B. thailandensis to utilize choline and its metabolites as carbon sources. Strains were grown with either 40 mM pyruvate (P), choline (C), GB (G), or DMG (D) as a sole carbon source in MOPS minimal media for

72 hours at 37 oC. (C) MBP-SouR binds to the putative promoter of the sarcosine oxidase operon represented by biotin-probe 3 in a dose dependent manner (lanes 3-5). The interaction is specific, as MBP-Sour does not bind to biotin-probe 2 (lane 8) and can be competed off by unlabeled probe 3 (lane 6 compared to lane 4, 22% shift versus 37.1% respectively), but not by unlabeled probe 2 (lane 7 compared to lane 4, 38.2% shift versus

37.1% respectively). Data in panels (A) & (B) are averaged from three experiments each with three biological replicates and error bars represent SEM. These data were both analyzed by two-way ANOVA with Tukey’s post-test testing for strain effect for each carbon source separately, not comparing the carbon sources. For (A), except for ΔgbdR2, all choline changes are different from WT and are not noted. Only the sarcosine comparisons different from WT are noted. For (B), pyruvates were not significantly different, but all other comparisons were except WT vs. the mutant in dimethylglycine. p

< 0.01 (**), p < 0.001 (***).

Figure 2.S1 – Complementation of ΔgbdR1ΔgbdR2 with gbdR1 or gbdR2.

Complementation of ΔgbdR1ΔgbdR2 with gbdR1 alone is able to restore the ability of B. thailandensis to utilize choline as a carbon source, but gbdR2 alone is unable to complement the metabolic deficiency. Data shown are averaged from three experiments each with three biological replicates and error bars represent SEM. Data analyzed by two-

66 way ANOVA with Dunnett’s post-test using the double deletion mutant as the comparator making comparisons within each time point. The asterisks indicate p < 0.01.

Figure 2.S2 – Validation of RNA-Seq experiments.

Validation of RNA-Seq experiments by qRT-PCR supports the findings of the RNA-Seq experiment. RNA-Seq values were normalized within DEseq based on library size while the RT-qPCR values were normalized to the rplU transcript levels. Both data sets are expressed here as fold change in expression scaled to their own respective y-axes.

Figure 2.S3 – pAN27 is still responsive to choline in ΔsouR. pAN27 is still responsive to choline in ΔsouR, and β-galactosidase activity seems to be induced to even higher levels as compared to wild type. This suggests that SouR is not required for the induction of glyA promoter. Data shown are averaged from three experiments each with three biological replicates and error bars represent SEM.

Figure 2.S4 – Usage of choline and sarcosine as sole carbon or nitrogen sources.

B. thailandensis can utilize choline as either a carbon or carbon and nitrogen source. B. thailandensis is unable to utilize sarcosine as a carbon or carbon and nitrogen source, unlike P. aeruginosa. Strains were grown in MOPS minimal media or MOPS minimal media without NH4, with either 40 mM or choline (C) or sarcosine (S) for 72 hours at 37 oC. Data shown are averaged from three experiments each with three biological replicates and error bars represent SEM.

2.8 Chapter 2 Tables

Table 2.1 – Transcript changes of B. thailandensis in response to choline.

Table 2.S1 – Strains and plasmids.

Table 2.S2 – Primers used in study.

Table 2.1 Transcript changes of B. thailandensis in response to choline.

Linear fold change in transcript over pyruvate

alonea:

Locus ID Geneb wt ΔgbdR1 ΔgbdR2 ΔgbdR1

ΔgbdR2

BTH_I0192 -2.6 NCc NC NC

BTH_I0393 -4.8 NC NC NC

BTH_I0394 -5.1 NC NC NC

BTH_I0395 -8.3 NC NC NC

BTH_I0396 -4.3 NC NC NC

BTH_I0398 -3.9 NC NC NC

BTH_I0687 23.8 NC 10.5 NC

BTH_I0688 19.3 NC NC NC

BTH_I0698 11.3 NC NC NC

BTH_I0700 12.0 NC NC NC

BTH_I0866 5.8 5.3 NC NC

BTH_I0959 -2.6 NC -5.3 -5.0

BTH_I1406 -2.3 NC -4.4 NC

BTH_I1620 5.7 NC 4.4 4.4

BTH_I1621 61.2 NC 18.3 NC

BTH_I1622 52.7 NC 9.7 NC

BTH_I1623 40.0 NC 9.3 NC

BTH_I1624 43.7 NC 5.0 NC

BTH_I1625 5.5 NC NC NC

BTH_I3016 -2.8 NC -3.6 -3.4

BTH_I3017 -3.1 NC -4.0 -3.9

BTH_II0001 5.4 NC 9.0 6.5

BTH_II0643 2.4 NC NC NC

BTH_II0694 -3.4 NC -2.4 NC

BTH_II0695 -2.6 NC NC NC

BTH_II0964 2.9 NC NC NC

BTH_II0966 cbcV 45.4 NC NC NC

BTH_II0967 cbcW 94.0 NC NC NC

BTH_II0968 gbdR2 81.7 NC NC NC

BTH_II0969 cbcX 136.6 NC NC NC

BTH_II0970 37.7 NC NC NC

BTH_II0971 8.8 NC NC NC

BTH_II0994 souR 3.9 NC 4.7 NC

BTH_II0995 sdaA-2 90.8 NC 32.3 NC

BTH_II0996 soxB 130.6 NC 39.3 NC

BTH_II0997 soxD 172.0 NC 102.3 NC

BTH_II0998 soxA 122.7 NC 58.6 NC

BTH_II0999 soxG 24.9 NC 11.7 NC

BTH_II1000 7.6 -3.2 8.7 -3.5

BTH_II1546 4.3 NC 3.7 NC

BTH_II1853 5.4 NC NC NC

BTH_II1854 cosX 7.1 NC NC NC

BTH_II1855 betC 3.8 NC NC NC

BTH_II1856 11.5 NC NC NC

BTH_II1857 purU2 8.4 NC NC NC

BTH_II1858 6.5 NC NC NC

BTH_II1859 betX 175.6 NC 38.6 NC

BTH_II1860 gbcB 115.8 NC 28.9 NC

BTH_II1861 gbcA 110.8 NC 23.7 NC

BTH_II1862 218.7 NC 44.4 NC

BTH_II1863 163.4 NC 97.2 NC

BTH_II1864 dgcB 154.1 NC 28.9 NC

BTH_II1865 dgcA 199.7 NC 23.4 NC

BTH_II1866 223.9 NC 29.8 NC

BTH_II1867 135.3 NC 47.4 NC

BTH_II1868 glyA 162.9 NC 20.7 NC

BTH_II1869 gbdR1 2.6 NC NC NC a Constraints on inclusion in table were +/- 2.3 fold change and p ≤ 0.005 based on the wild type. b Gene names are putative based on homology to P. aeruginosa. c NC, no change due to significance, lower fold change, or experimental deletion of the gene.

Table 2.S1 – Strains and plasmids.

Designation Genotype or Description Reference or Source

Burkholderia thailandensis strains MJ358 E264 wild type (23) AN3 ΔgbdR1 This study AN19 ΔgbdR2 This study AN16 ΔgbdR1 ΔgbdR2 This study AN126 ΔsouR This study AN202 ΔgbdR1 ΔsouR This study AN205 ΔgbdR2 ΔsouR This study AN207 ΔgbdR1 ΔgbdR2 ΔsouR This study AN136 attTn7 vector, Zeor This study AN153 attTn7 vector, Zeor This study AN128 attTn7 vector, Zeor This study AN150 attTn7 vector, Zeor This study AN158 ΔgbdR1 att::gbdR1 comp, Zeor This study AN173 ΔgbdR2 att::gbdR2 comp, Zeor This study AN162 ΔgbdR1 ΔgbdR2 att::gbdR1, Zeor This study AN164 ΔgbdR1 ΔgbdR2 att::gbdR2, Zeor This study AN151 att::souR comp, Zeor This study AN191 pAN27 This study AN192 ΔgbdR1 + pAN27 This study AN193 ΔgbdR2 + pAN27 This study AN194 ΔgbdR1 ΔgbdR2 + pAN27 This study AN192 ΔsouR + pAN27 This study AN226 pAN28 This study AN227 pAN29 This study AN228 pAN30 This study AN229 pAN31 This study

Escherichia coli strains DH5α T7 Express See manufacturer NEB AN133 pAN13 in T7 Express AN134 pAN14 in T7 Express

AN188 pAN19 in T7 Express

Plasmids pMQ132 Yeast recombineering vector, Gmr (38) pAN1 pMQ132 derivative, Tpr This study pAN1 derivative, lacZYA reporter pAN7 plasmid, Tpr This study pAN27 pAN7-PglyA -408 This study pAN28 pAN7-PglyA -340 This study pAN29 pAN7-PglyA -251 This study pAN30 pAN7-PglyA -147 This study pAN31 pAN7-PglyA -85 This study N-terminal MBP affinity tagging, pMAL-C2X Ampr NEB pAN13 pMAL-C2X-gbdR1 This study pAN14 pMAL-C2X-gbdR2 This study pAN19 pMAL-C2X-souR This study pUC18-mini-TN7T-Zeo attTn7 integration vector, Zeor (26) pTNS2 carrying the attTn7 transposase, Ampr (59) pAN23 pUC18-mini-TN7T-Zeo-gbdR1 comp This study pAN10 pUC18mini-TN7T-Zeo-gbdR2 comp This study pAN22 pUC18-mini-TN7T-Zeo-souR comp This study

Table 2.S2 – Primers used in study.

Bt-gbdR1- gcgccccggccgcatcaagatcagatccaaggaacgcttcGA gbdR1 (BTH_II1869) TpFLP-1F GCTCGAATTAGCTTCAAA deletion Bt-gbdR1- ggtgtgcgatttcgggggccggtttcgagttccggtttccGAGC gbdR1 (BTH_II1869) TpFLP-1R TCGAATTGGGGATCTT deletion Bt-gbdR1- CGTAATCGATCAGCATCGTG gbdR1 (BTH_II1869) 2F deletion Bt-gbdR1- GAAGCGTTCCTTGGATCTGA gbdR1 (BTH_II1869) 2R deletion Bt-gbdR1- GGAAACCGGAACTCGAAAC gbdR1 (BTH_II1869) 3F deletion Bt-gbdR1- ACGTCGAGACGCTGAGGAT gbdR1 (BTH_II1869) 3R deletion

BtgbdR2 AAG CCG TGC CGA TGT GGC TG gbdR2 (BTH_II0968) proximal F deletion BtgbdR2 AAG ATC GAC CAG CGG TAA TG gbdR2 (BTH_II0968) proximal R deletion BtgbdR2 tp ACTACGTCGCGCGCGCGGACCATTACCG gbdR2 (BTH_II0968) F CTGGTCGATCTTgagctcgaattagcttcaaa deletion REVISED BtgbdR2 tp TACGCCTTGCTGAAATGGCACGGCGAGT gbdR2 (BTH_II0968) R GAAAGCCGCAGAgagctcgaattggggatctt deletion REVISED BtgbdR2 TCTGCGGCTTTCACTCGCCG gbdR2 (BTH_II0968) post F deletion REVISED BtgbdR2 CGCCGCCGCTCAGATAGTCG gbdR2 (BTH_II0968) post R deletion REVISED BtII0994pr CGGCAATCTACGATGAGGCT souR (BTH_II0994) oxF deletion BtII0994pr CGATAGGGGGAAGAACACCG souR (BTH_II0994) oxR deletion BtII0994po GGTCAGCGTGAAGTTAGGCA souR (BTH_II0994) stF deletion BtII0994po CCGATGTCGTCCTTCTCGTC souR (BTH_II0994) stR deletion BtII0994tp GCCGCCGGCCGCATCGGCGTCGGTGTTC souR (BTH_II0994) F TTCCCCCTATCGgagctcgaattagcttcaaa deletion BtII0994tp GCGCTTCGGGATCGTGCTGCTGCCTAACT souR (BTH_II0994) R TCACGCTGACCgagctcgaattggggatctt deletion gbdR1com CCTAAGCTTGAGCGACATCCCGAGCACC gbdR1 complement pFHindIII G gbdR1com CTTGGTACCCGGGGGCCGGTTTCGAGTT gbdR1 complement pRKpnI C KpnIgbdR2 CTT GGTACC CTGACGTTTTCCGCCTTCG gbdR2 complement compF EcoRIgbdR CTT GAATTC GGGATCGGTTCGTGGCTG gbdR2 complement 2compR KpnI CTT GGTACC souR complement (use II0994com CCTTGCGTAACGATGCGTGGT MBP-SouR reverse pF primer) Tp TACGCCGTGGGTCGATGTTTGATGTTATG Generation of pAN1 pMQ132F GAGCAGCAACGgatcccctgattccctttgt Tp ACTCCGCGGCCGGGAAGCCGATCTCGGC Generation of pAN1 pMQ132R TTGAACGAATTGagcgcttttgaagctgatgt BtPFusion CTACTGCCGCCAGGCAAATTCTGTTTTAT Generation of pAN7 YHRp1F CAGACCGCTTC GAATTC CTGCCCG step 1 BtPFusion GCAGCTGGCACGACAGGTTTCCCGACTG Generation of pAN7

YHRp1R GAAAGCGGGCAG GAATTC GAAGCGG step 1 BtPFusion CTACTGCCGCCAGGCAAATTCTGTTTTAT Generation of pAN7 YHRp2F CAGACCGCTTC ccaagcttgcatgcctgcag step 2 BtPFusion GCAGCTGGCACGACAGGTTTCCCGACTG Generation of pAN7 YHRp2R GAAAGCGGGCAG step 2 gaattCGCAGCGTATCAGGC glyAtogbd CCTGCATGCAAGGCGTCTCGTCACTGGA Generation of pAN27 R1interF T glyAtogbd CTTGCATGCGGGGAAGCGTTCCTTGGAT Generation of pAN27 R1interR CT SphI CTT GCATGC Generation of pAN28, P.glyA1Lac CGCAACTAGCGTGCCATACG use with Z1 glyAtogbdR1interR SphI CTT GCATGC CTGCGCAGCGCGAGTG Generation of pAN29, P.glyA1Lac use with Z2 glyAtogbdR1interR SphI CTT GCATGC Generation of pAN30, P.glyA1Lac GAAAGCGTCTGAATTCATCAATCGG use with Z3 glyAtogbdR1interR SphI CTT GCATGC CCGCTGCGGCGGAAC Generation of pAN31, P.glyA1Lac use with Z4 glyAtogbdR1interR gbdR1EMS GGGGTTGGCGTTCGACATAA EMSA Probe 2 A2F gbdR1EMS CGGAAAAGGGAAAGAAAGCGT EMSA Probe 2 A2R BIOgbdR1 BIO-GGGGTTGGCGTTCGACATAA EMSA Probe 2 EMSA2F gbdR1EMS GCTAAACCCTAGATCGGCGG EMSA Probe 1 A3R gbdR2EMS CAGCGTATGCAGGAACGGG EMSA Probe 1 A1F BIOgbdR1 BIO-TGTCGTTCGGATTTTTAGCCG EMSA Probe 1 EMSA3F SarOxiEM CGGTCATGTCTGCAAAATCGT EMSA Probe 3 SA1F SarOxiEM GCCGATAGAACAAAAACGGCA EMSA Probe 3 SA1R BioSarOxi BIO-CGGTCATGTCTGCAAAATCGT EMSA Probe 3 EMSA1F pMALC2X CCTGGATCCTCGCCCGACCGCACCGCG MBP-GbdR1 gbdR1F RSFgbdR1 CTT AAG CTT TCA GCG CGC CGA CAC MBP-GbdR1 R GCG MALgbdR CCG GGATCC GTG ACG TCC GCC GCC MBP-GbdR2 2F GCT

6hisgbdR2 CTTAAGCTTTCAGCGCGCCGACACGCG MBP-GbdR2 R BamHI CCG GGATCC MBP-SouR II0994 GTGCGTTTCGGCGATGGTTC pMAL F HindIII CCT AAGCTT TCAGCGAGCGCCCCTC MBP-SouR and souR II0994 complementation pMAL R BtRplURT CAGTACAAGGTTGCCGTTGG rplU qRT-PCR set F BtRplURT CCGTGCTTTTGGTAGTGCTTC rplU qRT-PCR set R II1861RTF GCAGCAGATACGGCTCCATC gbcA qRT-PCR set II1861RTR GCAACGAGGAAAAAGGGTCG gbcA qRT-PCR set II0996RTF CTCGAACTACCTGTGGGACG soxB qRT-PCR set II0966RTR AAGTTGATCGTCGGCTCGAT soxB qRT-PCR set II0994RTF CGTTGCCTTTCTGCATCTCG souR qRT-PCR set II0994RTR TGCGTGAGCTGGTTCCATTA souR qRT-PCR set BtgbdR1 AGTGCTCAGGATGGCAAACT gbdR1 qRT-PCR set RT F BtgbdR1 CAGATTTTCCCAATGGATCG gbdR1 qRT-PCR set RT R gbdR2 set3 CAGGTGTCCGAGCAGTTCAT gbdR2 qRT-PCR set RTF gbdR2 set3 CGGCGACACGTTCAGATAGA gbdR2 qRT-PCR set RTR

2.9 Chapter 2 References

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CHAPTER 3:

EXPLORATION OF GATRS IN B. THAILANDENSIS

3.1 Abstract

Glutamine amidotransferase1 (GATase1)-containing AraC-family transcriptional regulators (GATRs) represent a widely disseminated class of regulator about which little is known. Given the particularly high presence of GATRs in β-proteobacteria, we choose to use the genetically tractable Burkholderia thailandensis as a model organism to increase our knowledge of these regulators. Phylogenetic comparisons of GATRs between Burkholderia species were made to determine how representative B. thailandensis is of other Burkholderiales. A small library of molecules was assembled based on functional predictions and potential orthologs of unknown GATRs. GATR deletions strains were screened against the library for growth on these compounds as either a carbon or nitrogen source, resulting in the identification of two additional GATRs in B. thailandensis.

To assess the functions of the remaining unknown GATRs, we developed a strategy to generate constitutive mutants by perturbing the predicted dimerization domain based on known constitutive mutations and a known crystal structure. We constructed a constitutively active mutant of a known GATR, GbdR, in P. aeruginosa, which led to the induction of genes involved in choline and glycine betaine transport and catabolism.

When this strategy was applied to a GATR of unknown function in B. thailandensis,

I0685, we found that I0685 acts as a repressor of an operon encoding glutathione-

91 dependent formaldehyde dehydrogenase. This function was additionally confirmed by growth assays in the presence of formaldehyde.

3.2 Introduction

Glutamine amidotransferase1 (GATase1)-containing AraC-family transcriptional regulators (GATRs) are a broadly occurring, yet poorly understood subfamily of AraC- family transcriptional regulators (ATRs). Currently there are few well-characterized members of the GATR subfamily. In order to better understand this subfamily of regulators, we sought to identify and characterize additional GATRs.

The ATR family is a prevalent and widespread family of regulators in bacteria (1-

3). ATRs are typically around 300 to 350 amino acids long. Structurally, ATRs may be divided into two domains separated by a short linker region (Figures 1.1). The C-terminal domain contains two helix-turn-helix motifs that are responsible for promoter recognition. The N-terminal domain is responsible for ligand binding and dimerization, and typically includes an unstructured N-terminal arm. ATRs typically function as dimers and bind two degenerate direct repeat half sites proximal to the transcriptional start site where they function as activators. The mechanism of activation for ATRs involves the perturbation of dimerization between monomers caused by ligand binding.

The namesake member of the family, AraC, activates transcription by the ‘light- switch’ mechanism whereupon transcription is repressed by DNA looping due to a preference for a distal upstream half site (Figures 1.2) (4). Binding of arabinose to the

AraC dimer results in a shift in preference for binding sites to the two half sites proximal

92 to the transcriptional start site, thereby activating expression of the target operon. This shift in orientation of the dimer is caused by interaction between the N-terminal arm, and the ligand binding domain when arabinose is bound (5). Alternatively, dimers of other

ATRs such as RegA remain in solution until their ligand is bound, which then causes the nature of the dimer to change and bind to its target promoter and activate transcription

(Figures 1.3) (3, 6).

GATRs are distinguishable from other ATRs based on the homology of the N- terminal dimerization and ligand recognition domain to the GATase1 family of enzymes

(7, 8). GATases catalyze the transfer of amide groups to various substrates (9). GATase1 domain-containing enzymes include anthranilate synthases, CTP synthases, GMP synthetases, and 2-amino-2-desoxyisochorismate synthases among many more, which are responsible for producing a wide range of both primary and secondary metabolites (10-

12). GATase1 domains may be fused to a synthase domain or translated as an independent peptide, which forms a heterodimer with a synthase domain (9). GATase1 domain-containing proteins often function as dimers or sometimes higher order oligomers. In some cases, higher order oligomerization leads to the production of proteinaceous filaments, such as in CTP synthetase, where filament formation negatively regulates enzyme activity in the presence of excess product (13). Many GATRs also share a conserved cysteine, the remnant of a cysteine/glutamate/histidine catalytic triad stemming from the homology to GATase1 enzymes.

GATRs are found in both Gram-positive and Gram-negative bacteria, with a notably higher presence in proteobacteria based on currently available sequence data

(14). While many GATR-possessing bacteria encode only one or two homologs, other

93 genomes may contain more. For example, Pseudomonas aeruginosa encodes seven (15) and Burkholderia cenocepacia encodes fifteen (16). To investigate the diversity and functionality of GATRs, Burkholderia thailandensis was chosen as a model organism based on the availability of genetic tools and its genetic tractability (17-20).

The genome of B. thailandensis contains thirteen bioinformatically predicted

GATRs including putative orthologs of all GATRs present in the closely related select agents Burkholderia pseudomallei and Burkholderia mallei, the etiological agents of melioidosis and glanders (21, 22). B. thailandensis also contains putative orthologs of

GATRs found elsewhere in the Burkholderia genus, making the model a respectable survey of the GATR family found here and in more distally related bacteria (Figures 3.1,

3.2, and Table 3.4).

Previously identified GATRs in P. aeruginosa, GbdR and ArgR, have been shown to function as transcriptional activators for genes involved in the transport and catabolism of glycine betaine (GB) and arginine respectively (23-26). To better understand the function of B. thailandensis GATRs, and what compounds GATRs might enable the bacterium to utilize, we generated deletion strains of each GATR.

Synthesizing the functional predictions for adjacent and co-occurring genes with our knowledge of known GATRs, deletion strains were screened against a collection of potential metabolites as sole carbon and nitrogen sources. Using these screen we identified orthologs of the GATRs ArgR and CdhR.

In order to identify the regulons of additional unknown GATRs, we designed an approach based on perturbation of the dimerization of GATRs to achieve differential expression of their target operons independent of ligand by combining information about

94 known constitutive mutants in characterized GATRs with predicted protein structures.

Utilizing this method, we found that the unknown GATR, I0685, is responsible for regulating glutathione-dependent formaldehyde detoxification. Interestingly, this regulation is due to repression by I0685 revealing that GATRs may act as repressors, contrary to previous data that suggested GATRs function solely as activators (26-29).

The regulation of formaldehyde metabolism is important for bacteria, as formaldehyde at high concentrations may damage and interfere with other biological processes (30). Simultaneously, lower amounts of formaldehyde may be useful as singly carbon building blocks for more complex molecules (31). Choline catabolism in B. thailandensis produces three molecules of formaldehyde per molecule of choline, and three different GATRs are already known to impact the regulation of this pathway (32).

Formaldehyde can also be generated in bacteria as a result of methylglyoxal degradation, which can be generated by glyceraldehyde oxidation and the degradation of glyceraldehyde-3-phosphate or dihydroxyacetone-phosphate (33). The finding of an additional GATR that impacts formaldehyde catabolism is an interesting link to the

GATR-regulated catabolism of choline.

3.3 Results

3.3.1 Phylogenetic comparison of GATRs in Burkholderia.

Alignment of the full length sequences of Burkholderia species GATRs resulted in a general grouping of the GATRs into three groups (Figure 3.1). The first group consists of orthologs of GbdR1, GbdR2, SouR, CdhR, and ArgR, all except ArgR

95 regulate different metabolic pathways related to GB catabolism. The second group includes I1350, I0685, II1912, II0513, and several B. cenocepacia GATRs that do not have apparent orthologs in the other Burkholderiales. The third group includes II2197,

II0282, and II1760. All the members of this clade are bioinformatically labeled as FtrA or

FtrA-like, a GATR accociated with accessory nitrogen metabolism present in

Caulobacter crescentus (34). However, upon comparison of the clustering with FtrA from Yersinia enterocolitica and C. crescentus, and the conserved synteny of a putative rhodanese divergently transcribed from it, II1760 is likely the genuine ortholog of FtrA as identified from E. coli (Figures 3.1, 3.2). Because the DNA binding domain might drive alignments due to similarities in HTH motifs subjected to possible selective pressure based on similarities between target binding sites, we also generated a phylogenetic tree based on alignment of only the GATase1-like domain (Figure 3.2). The similar results suggest that the alignments produced were not strongly influenced by the C-terminal domains. The GATR I0059 does not fit into any one of the clade in either alignment.

To further investigate the groupings of putative orthologs derived from the alignments, gene synteny and co-occurrence of the nearby genes using the STRING database and the Burkholderia Genome Database were utilized (16, 35), generating Table

3.4. These comparisons suggest that the phylogenetic clustering is accurate. However, the association of BCAS0761 with a particular group based on Table 3.4 was not clear. The phylogenetic alignments group BCAS0761 with I0685 and it co-occurs with orthologs of genes encoding glutathione-dependent formaldehyde dehydrogenase and an associated esterase (I0686 and I0687). Orthologs of I0686 and I0687 in P. aeruginosa are regulated by GfnR, a LysR-family regulator. GfnR is not present in B. cenocepacia suggesting an

96 alternative regulator such as an ortholog of I0685 may be required for regulation of these genes (28).

B. mallei does not contain an ortholog for SouR, despite possessing orthologs for

GbdR1, GbdR2, and CdhR.

3.3.2 GATR functionality by screening carbon and nitrogen utilization.

Our knowledge of the inducing ligands for GbdR and ArgR in P. aeruginosa (25,

36) and bioinformatic predictions for genes either adjacent or co-occurring with GATRs

(Figure 1.7), allowed for the assembly of a collection of compounds to screen as a sole carbon or nitrogen sources to determine if the GATR deletions displayed phenotypes under any of these conditions. Deletion strains were generated for every GATR in B. thailandensis, except for II1912, which differs in length and orientation between differing strains of B. thailandensis, and may not conserved or functional in B. thailandensis E264.

ΔgbdR1, ΔgbdR2, and ΔsouR strains exhibited growth defects on choline, glycine betaine, dimethylglycine, and sarcosine as sole carbon sources (Figure 3.3). ΔgbdR1 and

ΔgbdR2 grew on sarcosine as a sole nitrogen source. However, the ΔsouR strain demonstrated no growth on sarcosine (Figure 3.4). These results support our previous findings (32), thus serving as effective controls for the screen. Different strains appeared to exhibit large differences in growth rates on a large set of compounds, which may be partially explained by differences associated with the biofilm phenotype discussed in

Appendix 1.

ΔgbdR1, ΔgbdR2, and ΔsouR strains were unable to utilize choline-O-sulfate as

97 either a sole carbon or nitrogen source compared to wild type (Figures 3.3 and 3.4).

Choline-O-sulfate is utilized in some plants, fungi, and bacteria as a compatible solute or as management for high levels of sulfate (37-39).

Two other deletion strains exhibited growth defects in the screen. ΔII1850 was unable to grow on carnitine as either a sole carbon or nitrogen source (Figures 3.3 and

3.4). This suggests that II1850 is likely an ortholog for CdhR, a GATR in P. aeruginosa shown to regulate genes for carnitine transport and catabolism (40). The second strain,

ΔI1774, was defective in growth on arginine as a sole carbon source (Figure 3.3) and appeared to utilize arginine as a sole nitrogen source at a diminished rate compared to wild type and other strains (Figure 3.4). This result suggested that I1774 may be an ortholog for ArgR, a GATR in P. aeruginosa responsible for regulating genes involved in arginine synthesis and catabolism (26).

3.3.3 Deletion of II1850 results in a catabolic defect on carnitine as a sole carbon source.

The synteny of genes adjacent to cdhR in P. aeruginosa and II1850 are identical and encode similar proteins, suggesting II1850 is a cdhR ortholog (Figure 3.5A). The cdhA, cdhB, and cdhC genes encode proteins directly responsible for the catabolic degradation of carnitine into glycine betaine (40). caiX, a gene encoding a periplasmic binding protein, is responsible for the transport of carnitine from the periplasmic space through interaction with CbcWV, an ATP-binding cassette transporter (41). Finally, hocS

98 encodes hydrolase responsible for the degradation of short-chain O-acetyl carnitines that also feeds into this catabolic pathway (42).

Growth of ΔII1850 on carnitine as a sole carbon source on MOPS minimal media plates confirms the catabolic defect previously observed in catabolic screens (Figure

3.5B). This defect was complemented by addition of II1850 back onto the chromosome at the attTn7 site, under the control of its native promoter (Figure 3.5B, right panel). The wild type, deletion, and complement strains are all capable of growing on pyruvate under these growth conditions (Figure 3.5B, left panel).

3.3.4 Deletion of I1774 results in a catabolic defect on arginine as a carbon source.

Comparison of the genomic context of I1774 in B. thailandensis with the genomic context of argR in P. aeruginosa reveals an adjacent operon containing predicted arginine succinyltransferase pathway (AST) genes (Figure 3.6A). These GATRs are both located at the end of a transport operon. This operon is described as being responsible for the transport of arginine and ornithine under the regulatory control of ArgR (25). The arrangement of these operons differs between these species, with the B. thailandensis operon containing only putative orthologs to aotQ (I1771), aotM (I1772), and aotP

(I1773). A potential ortholog for aotJ (I1783), a periplasmic binding protein that primarily contributes to the uptake of arginine (25), is located proximally to AST genes.

Additionally, an ortholog for autO (II2133) exists, but it is not strictly required for arginine catabolism (25).

Deletion of I1774 results in a defect in growth on arginine as a sole carbon source

(Figure 3.6B, right panel). This defect is complemented by addition of the I1771-4 operon at the attTn7 site under the control of its native promoter. All strains were able to grow on pyruvate as a sole carbon source under these conditions (Figure 3.6B, left panel).

Efforts to complement the phenotype using just I1774 and the immediate 200 bp upstream from it were unsuccessful, likely because I1774 is part of an operon and the native promoter required for expression is located upstream from I1771 (data not shown).

3.3.5 GbdR E36G constitutively upregulates transcription of members of the GbdR regulon.

Discovering the function of the remaining unknown GATRs presented a hurdle as neither their ligand nor the genes they regulate are known. The construction of constitutively active transcriptional regulators might be used to identify genes that are upregulated in a ligand independent manner. In order to determine whether constitutively active GATRs can be constructed based on examples of constitutive mutation in SouR from P. aeruginosa and the crystal structure 3GRA from P. putida (Figure 3.7), we focused on GbdR in P. aeruginosa as a model since its regulon and function is understood (23). A GbdR E36G strain was designed based on the predicted structural homology to the D48G allele constitutive mutant of SouR in P. aeruginosa PA14 (Table

3.5). This mutant allele was capable of rescuing the catabolic phenotype of ΔgbdR on choline as a sole carbon source (Figure 3.8), demonstrating that the protein is functional.

When grown in the absence of choline, the inducing ligand, microarray analysis revealed

100 that transcripts in the GbdR regulon were upregulated (Table 3.6). These include genes responsible for the transport of choline and its metabolites such as cbcW and betX, as well as genes encoding enzymes that catabolize GB into glycine such as gbcA, soxA, soxB, and glyA among others. This result suggests successful construction of a constitutively active, ligand-independent version of GbdR. Similar mutations and analyses can be used to uncover the regulons of other GATRs.

3.3.6 I0685 regulates expression of a formaldehyde dehydrogenase.

Similarly to our approach with GbdR, we constructed a ligand independent, constitutive I0685 by incorporating an N32G mutation based on comparison to constitutive mutations in SouR of P. aeruginosa and the crystal structure of 3GRA in P. putida (Table 3.5). I0685 is divergently transcribed from a highly conserved operon that includes a glutathione-dependent formaldehyde dehydrogenase and an associated esterase. Surprisingly, when the transcriptome of the N32G expressing strain was compared to the deletion strain, the adjacent dehydrogenase and esterase genes were significantly downregulated in the presence of the predicted “constitutive” mutant (Table

3.7). This was opposite of the result we expected to observe based on our knowledge that other GATRs function as activators (24, 26). The downregulation is dependent on the presence of the N32G mutant, and not simply due to accidental disruption of the promoter during deletion of I0685. It is possible that the mutation has resulted in a conformational change that causes insensitivity to de-repressing ligand that might normally be present.

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In order to further assess the potential impact on formaldehyde detoxification,

I0685 deletions and complement strains were constructed in a ΔII1870 (orthologs commonly named fdhA) background in order to separate the impact of the glutathione- dependent formaldehyde dehydrogenase from redundant detoxification by the glutathione-independent formaldehyde dehydrogenase. Previously, the multiple systems present in Burkholderia have been shown to successfully compensate for each other (43).

Growth in the presence of formaldehyde revealed that the I0685 deletion had a slight growth advantage compared to the wild type. Complementation of I0685 back onto the chromosome was slightly deleterious, and complementation with the N32G mutant greatly inhibited growth in the presence of formaldehyde (Figure 3.9). This suggested that the I0685 may function to repress the expression of I0686 and I0687.

3.4 Discussion

3.4.1 Building constitutive regulators.

The construction of constitutively active transcriptional regulators is a useful tool both for uncovering details of mechanism and, as we have explored here, revealing target genes. Constitutive mutants in AraC have been previously published, with the majority being found to be localized in the N-terminal arm (44). Other constitutive mutations were found to occur in the dimerization domain itself and some in the DNA binding domain

(45). Contrary to the experience of other groups, all constitutive mutations we have identified experimentally occur in the predicted dimerization face of the N-terminal domain (Figure 3.7). Although GATRs also appear to possess an unstructured N-terminal

102 arm, it is not well-conserved and it is unclear if the N-terminal arm plays an important mechanistic role in GATR function. However, GATase functionality is heavily dependent on ligand induced conformational changes, which are necessary for proper formation of ammonia channels to supply associated synthase domains (8). Because dimerization or higher oligomerization of GATase-containing enzymes is critical to proper formation of ammonia channels and orientation of active sites for functionality, it stands to reason that altering the ability to undergo conformational changes through mutations in the dimerization domain might greatly affect the functionality of the protein. Indeed, constitutive GATases have been constructed through mutation of residues critical for proper active conformation (46). It is possible that the homology of GATRs to GATase1 enzymes have contributed a mechanism for conformational change allowing GATRs to function through a conformational change-based mechanism similar to the mechanisms of other ATRs such as AraC and RegA.

In order to test our ability to generate constitutive GATRs, we utilized GbdR from

P. aeruginosa as a model. GbdR is responsible for regulation of enzymes in the GB catabolic pathway, various transporters, and the extracellular phospholipase PlcH (23, 24,

27). This pathway converts choline into the ubiquitous osmoprotectant GB and can further catabolize it through a series of demethylation steps ultimately into glycine. B. thailandensis also shares this catabolic pathway, although the gene organization and regulation scheme differs from P. aeruginosa and includes two GbdR orthologs instead of just one (32). GbdR is thought to bind DNA as a dimer due not only to similarities to

ATRs, but more specifically to ArgR which has been shown to dimerize and share similar structure of the orientation of the direct repeat binding sites of within target promoters

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(23, 25, 47). Using the combined information from the crystal structure of the N-terminal domain of 3GRA from P. putida and the constitutive mutants of SouR from P. aeruginosa, we were able to successfully generate a constitutive GbdR that activates transcription of target genes by altering a residue in the predicted dimerization domain.

The alteration to the dimerization domain may alter the conformational preference of the dimer, resulting in a bias toward the activating conformation even in the absence of ligand. Currently it is unclear whether the change in conformation of this constitutively activating GbdR is altering an interaction with the RNA polymerase holoenzyme or changing the way in which GbdR acts with its DNA binding half sites, as in the case for

AraC.

3.4.2 Identification of a GATR acting as a repressor of formaldehyde detoxification.

Formaldehyde is a biologically significant molecule because it can be a useful one carbon building block, or a highly reactive molecule damaging proteins and DNA.

Accordingly, the metabolism of formaldehyde must be tightly controlled (33).

Consequently, bacteria utilize several pathways for formaldehyde detoxification. B. thailandensis is predicted to contain two formaldehyde detoxification pathways: a glutathione-independent formaldehyde dehydrogenase and a glutathione-dependent formaldehyde dehydrogenase. Other Burkholderiales have been shown to contain other, alternative pathways for formaldehyde detoxification. Burkholderia cepecia TM1 possesses genes encoding a ribulose monophosphate-dependent pathway shown to be important for the detoxification of formaldehyde generated by the degradation of lignin from plant matter (48, 49). Closely related and previously classified as a Burkholderia,

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Paraburkholderia fungorum LB400 utilizes the pterin-dependent formaldehyde detoxification pathway which is dependent on either the pterin-containing tetrahydrofolate or tetrahydromethanopterin. This pathway is widespread amongst bacteria due to utilization in the one carbon transfer needed for the synthesis of purines and amino acids (33, 43).

The glutathione-independent formaldehyde dehydrogenase, II1870, is adjacent to gene required for GB catabolism. Transcription of II1870 may be under the regulatory control of the GbdR orthologs GbdR1 and GbdR2 in addition to a conserved raffinose repressor in the same operon (32). This reaction relies on NAD+ and results in the generation of formate (50). The close association between II1870 and GB catabolism is likely due to the three molecules of formaldehyde produced for every molecule of GB that is catabolized to pyruvate (36).

The glutathione-dependent pathway utilizes glutathione to react with formaldehyde resulting in S-hydroxymethylgluathione, which is then converted to S- formylglutathione by the formaldehyde dehydrogenase (I0686 in B. thailandensis) (51,

52). An associated esterase (I0687 in B. thailandensis) then converts this product into formate and regenerates glutathione (53). While the glutathione-dependent formaldehyde dehydrogenase and esterase are regulated by the LysR-family transcriptional regulator

GfnR in P. aeruginosa and many other bacteria, B. thailandensis has no ortholog for

GfnR. Instead an unknown GATR, I0685, is divergently transcribed from the operon containing these genes (28). Operons divergently transcribed from their regulator is a common gene arrangement (54).

We applied our strategy for constructing constitutive GATRs to I0685. We

105 expected a ligand-independent upregulation of the glutathione-dependent formaldehyde dehydrogenase and esterase genes. Surprisingly, the deletion strain expressing I0685

N32G significantly downregulated expression of these genes when compared to an I0685 deletion strain carrying the empty vector (Table 3.7). The change in expression of these genes stimulated the investigation of the effects of formaldehyde on the growth of various strains. Strains producing wild type, or particularly I0685 N32G, have reduced tolerance of formaldehyde compared to the deletion itself, likely due to direct repression of I0686 and I0687.

The I0685 regulatory mechanism is novel in multiple ways. First, characterized

ATRs are typically activators (3, 55) and this marks the first incidence of any GATR acting specifically as a repressor. Second, GATRs have been associated with smaller amine-containing compounds such as arginine, carnitine, GB, and sarcosine. However,

I0685-7 are not transcribed in a GB- or GbdR- dependent manner, and the upregulation of I0686-7 has been observed in connection with the catabolism of GB, which produced formaldehyde (32). It is quite possible that formaldehyde itself is the ligand to which

I0685 is responding, which suggests not all GATRs respond to amine-containing compounds as we had initially suspected.

3.4.3 II1850 regulates carnitine catabolism.

Carnitine is an abundant metabolite in many environments (56) and found in particularly high levels in muscle tissue (57). Carnitine is also found in plants, but in far lower concentration than in animals (58). Animals use carnitine to shuttle fatty acids by

106 using O-acylcarnitine shuttles, with the acylcarnitines being generated by the liver (59-

61). Interest in carnitine has been piqued recently because gut microbiota have been shown to metabolize dietary carnitine into trimethylamine that is then converted by the host into trimethylamine-N-oxide, which has been linked to atherosclerosis (62, 63). In addition to utilizing carnitine as a carbon and/or nitrogen source, examples of both Gram- negative and -positive bacteria have been shown to use carnitine as an osmoprotectant directly, or by catabolism into GB (40, 64-67).

II1850 is clearly an ortholog of CdhR in B. thailandensis given the similarity in gene synteny and phenotype of the deletion strains when compared to P. aeruginosa

(Figure 3.5). The presence of a carnitine catabolic pathway provides another route to acquire the osmolyte GB. In addition, the pathway provides yet another opportunity for taking advantage of available nutrients that may be present when the bacterium is in association with decaying plant or animal matter in the environment.

3.4.4 I1774 regulates arginine catabolism.

The metabolism of arginine in Gram-negative bacteria is well characterized and typically regulated in large part by one of two unrelated regulators named ArgR. ArgR in

E. coli is a hexameric repressor regulating genes involved in arginine transport and metabolism (68, 69). In bacteria lacking the hexameric repressor, a GATR alternative

(also called ArgR) is often found. The GATR ArgR has been shown to regulate genes involved in arginine biosynthesis, as well as genes in the AST pathway in P. aeruginosa

(26). This pathway is one of four pathways responsible for arginine utilization and is

107 considered the primary pathway responsible for the aerobic catabolism of arginine (70).

ArgR in P. aeruginosa regulates ldcA, a gene responsible for lysine catabolism, in an arginine dependent manner. It is hypothesized that the lack of lysine-responsive control of these genes accounts for poor utilization of lysine as a sole carbon source, despite being able to utilize lysine when arginine is present (71). A similar regulation scheme may be present in B. thailandensis, either due to regulation of lysine decarboxylases or the specific regulation and specificity of the assorted substrate periplasmic binding proteins. The presence of such a regulation scheme might explain the lack of growth of B. thailandensis strains on lysine as a sole carbon source in our screen

(Figure 3.3).

Based on the catabolic phenotype and high degree of homology between parts of the arginine transport and catabolic pathway components, I1774 is an ortholog of ArgR.

Similarities between components of the P. aeruginosa AST pathway and observed phenotypes in B. thailandensis suggest a similar approach to regulation in both organisms.

3.4.5 II1760, a potential regulator of cyanide detoxification.

Many Proteobacteria possess a single GATR that has been bioinformatically classified as FtrA, while B. thailandensis contains three GATRs that are currently annotated as being FtrA-like: II0282, II1760, and II2197. In our phylogenetic analysis of the GATRs (Figure 3.11A), we included typical examples of FtrA from E. coli, C. crescentus, and Y. enterocolitica. The resulting phylogenetic trees definitively clustered these GATRs with II1760 and putative homologs from other Burkholderia. The synteny

108 of the divergently transcribed gene, II1761, versus its putative homologs is conserved in all of these examples (Figure 3.11B). This suggests that this adjacent gene is a likely regulatory target for II1760.

II1761 is predicted to encode rhodanese, an enzyme that detoxifies cyanide by converting it to thiocyanate (Figure 3.11C) (72). The toxicity of cyanide is often due to inhibition of cytochrome c or other electron receptors, consequently impairing aerobic respiration (73). Rhodanese-like proteins are widely distributed among eukaryotes, prokaryotes, and archaea, and may also contribute to sulfur and selenium metabolism in addition to their role in cyanide detoxification (74). Many bacteria, specifically those with rhodanese, are of interest as potential bioremediation agents (75, 76). B. cepacia has been previously shown to degrade a wide range of cyanide containing compounds (77).

Bacteria and fungi often produce cyanide containing compounds to facilitate competition or avoid predation (78). An antagonistic relationship between P. aeruginosa and B. cenocepacia has also been described in the cystic fibrosis lung environment wherein cyanide containing compounds produced by P. aeruginosa inhibit the growth of a particular strain of B. cenocepacia (79).

This raises the possibility of a GATR responsible for cyanide detoxification and that potentially uses cyanide as a ligand. More direct evidence of regulation of rhodanese by II1760 in B. thailandensis is still needed, and the function of this putative rhodanese remains to be ascertained. It is likely that this information will reflect the function of

FtrA, perhaps the most pervasive GATR in proteobacteria, and its relationship to rhodanese in other species.

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3.4.6 Significance of choline-O-sulfate.

Some plants rely on osmoprotectant other than glycine betaine. In particular the

Plumbaginaceae, a salt-tolerant group of plants, have evolved to accumulate choline-O- sulfate to high levels in contrast to osmoprotectants such as glycine betaine (80). Choline-

O-sulfate has also been speculated to be involved in bacteria-plant communication (81).

B. thailandensis possesses genes predicted to transport and catabolize choline-O-sulfate, which were found to be induced in the presence of choline in our previous studies (32), and could be expected to feed into the downstream choline and glycine betaine catabolic pathways. Because the precise environmental niche of many Burkholderia species is not yet fully appreciated, perhaps this metabolic capacity is suggestive of co-occurrence with organisms commonly capable of producing and exuding choline-O-sulfate.

3.4.7 The absence of SouR in B. mallei.

Although it is tempting to speculate that one of the remaining GATRs may be responsible for regulation of the downstream portion of GB catabolism, further examination reveals that there are no putative orthologs of tetrameric sarcosine oxidase by BLAST (32). Sarcosine may also be catabolized by monomeric sarcosine oxidase (82,

83), but a putative ortholog is not obvious. It is possible that B. mallei either lacks the ability to utilize GB metabolites as carbon or nitrogen sources or that it possesses an alternative pathway not yet appreciated.

3.5 Methods

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3.5.1 Alignment of GATRs and phylogenetic tree construction.

The full length sequence of all GATRs from B. thailandensis E264, B. pseudomallei K96243, B. mallei ATCC 23344, B. cenocepacia J2315, and several

GATRs bioinformatically identified as orthologs of FtrA, in addition to AraC and PyrG from Escherichia coli, were aligned using the MUSCLE algorithm (84). In addition, the predicted GATase1-like domains for these GATRs and PyrG were also aligned and compared in order to eliminate similarity driven by HTH domains that bind similar DNA sequences. Maximum likelihood phylogenetic trees were generated using the MEGA6 software package (85). Each phylogenetic tree was bootstrapped using five hundred bootstrap repetitions.

3.5.2 Culture conditions for strains.

B. thailandensis, P. aeruginosa, and E. coli strains were typically plated on

Lennox broth (LB) (10 g/L tryptone, 5 g/L yeast extract, 5 g/L NaCl, 1.5% agar) agar and grown at 37oC prior to generating overnights for experiments. Modified MOPS minimal

+ media (86, 87) with and without NH4 was used for growth on particular carbon or nitrogen sources, as well as inductions.

3.5.3 Construction of deletion strains.

Deletion strains in B. thailandensis were generated by inducing natural competency in combination with the introduction of splice overlap extension PCR products as described in Thongdee et al. using primers in (Table 3.3) (17). Trimethoprim resistance markers in

111 the resulting insertion-deletion mutants were removed utilizing pFLPe4 as described by

Choi et al. (18).

3.5.4 Carbon and nitrogen utilization screens.

A custom selection of compounds was derived from bioinformatics, functional predictions based on gene synteny, and co-occurrence of genes associated with unknown

GATRs using the STRING database (35). Compounds for the carbon-source screen included pyruvate as a positive control, arginine, asparagine, β-alanine, N,N-dimethyl-β- alanine, 4-aminobenzoic acid (PABA), glutamate, glutamine, glycine, histidine, proline, hydroxyproline, serine, O-acetyl L-serine, ornithine, lysine, threonine, taurine, histamine, ethanolamine, agmatine, canavanine, citrulline, ecotine, theanine, xanthine, urea, uric acid, trigonelline, ɣ-amino butyric acid (GABA), ɣ-butyrobetaine (GBB), guanidine, aminoethylethanolamine (AEEA), carnitine, choline-O-sulfate, choline, glycine betaine

(GB), dimethylglycine (DMG), sarcosine, cadaverine, spermidine, putricine, N-acetyl-D- glucose, N-acetyl-D-galactose, UDP-glucose, creatine, and creatinine. The nitrogen- source screen included arginine, asparagine, β-alanine, N,N-dimethyl-β-alanine, cysteine, glutamate, glutamine, glycine, histidine, hydroxyproline, hydroxyserine, lysine, O-acetyl-

L-serine, ornithine, phenylalanine, proline, serine, threonine, tryptophan, taurine, agmatine, canavanine, citruline, ectoine, thenine, xanthine, urea, uric acid, trigonelline,

GBB, GABA, creatine, creatinine, carnitine, choline-O-sulfate, choline, GB, DMG, sarcosine, cadaverine, spermidine, putrescine, UDP-glucose, N-acetyl-glucosamine, N- acetyl-galactosamine, anthranilate, and PABA. Compounds were prepared in 48-well plates (Costar), one compound per well. For carbon-source screen plates, compounds

112 were added at 100 mM in 30 µL for a final concentration of 20 mM. For nitrogen-source screen plates, compounds were added at 25 mM in 30 µL for a final concentration of 5 mM. WT and deletion strains were struck onto LB agar plates and incubated overnight at

37oC. These were used to inoculate 3 mL of MOPS with 20 mM pyruvate and 5 mM glucose, which were incubated at 37oC overnight on a culture wheel. For carbon source screening, overnights were collected by centrifugation, washed with 1 mL MOPS, and resuspended in MOPS to an OD600 measurement of 0.05. For nitrogen source screening,

+ overnights were spun down and washed with 1 mL MOPS minus NH4 plus 20 mM

+ pyruvate, then resuspended in MOPS minus NH4 plus 20 mM pyruvate to an OD600 measurement of 0.05. In both cases, 120 µL of resuspended cells were added per well of the appropriate plates, which were sealed with sterile, gas-permeable plate film. The

o plates were incubated at 37 C in humidity chambers on a shaker, and OD600 was measured at 24 and 48 hours.

3.5.5 Prediction of amino acid mutations resulting in constitutively active GATRs.

Understanding of the putative dimerization of GATRs is based off of the crystal structure of the N-terminal domain of the SouR ortholog, 3GRA, originating from

Pseudomonas putida (88, 89). Unpublished data from Graham Willsey at the University of Vermont indicated mutations in particular residues of P. aeruginosa SouR that resulted in constitutive activation of SouR-regulated genes (Figure 3.7). The N-terminal domain of P. aeruginosa SouR was strung onto the 3GRA crystal structure using UCSF Chimera

(90), which revealed that the constitutive mutations were localized in the putative

113 dimerization domain. Portions of the N-terminal domain from various GATRs were then modeled onto the structure of 3GRA to identify the structurally orthologous residues in each GATR, resulting in the predictions contained within Table 3.5.

3.5.6 Generation of complement and constitutive GATR strains.

Plasmid tools for attTn7 integration produced by Choi and colleagues (19, 20, 91) were used for complementation in both B. thailandensis and P. aeruginosa. For B. thailandensis, complementing plasmids based on pUC18T-mini-TN7T-TP or pUC18- mini-TN7T-Zeo (Table 3.2) were co-transformed with pTNS2 using electroporation and subsequently selected on either LB agar plates containing 100 µg/mL trimethoprim or

Low-salt LB plates (3 g/L NaCl, pH= 7.5 with NaOH) containing 250 µg/mL zeocin. For

P. aeruginosa, a tri-parental mating strategy was utilized. Briefly, WT or ΔgbdR P. aeruginosa PA14 was combined with E. coli S17-1 containing pUC18-mini-TN7T-Gm based complement or empty vector and E. coli S17-1 containing pTNS2 in various proportions. Prior to conjugation, P. aeruginosa strains were heat-shocked for fifteen minutes at 42oC. These mixtures were spotted onto LB plates without antibiotics and incubated overnight at 30oC. Spots were scrapped off the plate and re-plated onto

Pseudomonas isolation agar (PIA) (BD) plates containing 50 µg/mL gentamicin and incubated overnight at 37oC. Resistant colonies were subcultured on PIA with 50 µg/mL gentamicin and selected clones were screened by PCR.

3.5.7 Growth of strains and RNA extractions.

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Putative constitutive complement mutants were grown in tandem with their respective deletion strain as a control to avoid potential interference by wild type GATRs either through interaction between mutant and wild-type GATRs or competition for DNA binding sites. 3 mL MOPS with 20 mM pyruvate and 5 mM glucose cultures were inoculated from LB plates and grown on a rotary wheel at 37oC. These overnights were pelleted in a table top centrifuge and washed once in 1 mL MOPS media with 20 mM pyruvate and resuspended in 1 mL of the same. OD600 measurements were taken so that induction cultures could be inoculated at a starting OD600 of 0.3, and these were grown for 4 hours at 37oC on a rotary wheel. Inductions were pelleted and resuspended in 800

µL of 95oC RNAzol RT (Sigma) whereupon they were frozen at -80oC after five minutes at room temperature.

To continue the RNA extraction, samples were thawed and 400 µL water was added to each sample and vortexed vigorously, followed by a room temperature incubation for fifteen minutes. Samples were centrifuged at 10,000 x g for fifteen minutes at 4oC, and the supernatant was transferred to a clean microcentrifuge tube. The supernatant was then combined with 100% molecular grade ethanol at a 1:1 ratio, mixed, and applied to Zymo-Spin IIC columns (Zymo Research (ZR)). Columns were centrifuged for thirty seconds at 10,000 rpm, washed with 400 µL direct-zol prewash

(ZR), then washed with 700 µL and 400 µL of wash buffer (ZR). The columns were further centrifuged to dryness for two minutes at 10,000 rpm, then RNA was eluted using

100 µL RNase-free water into a clean microcentrifuge tube.

In order to remove DNA, 11 µL of 10x DNase I Buffer and 1 µL of RNase-free

DNase I (New England BioLabs) were added to each sample and incubated at 37oC for

115 twenty minutes. An additional 1 µL of RNase-free DNase I was added to each sample and the incubation at 37oC was continued for another twenty minutes. Samples were then further processed using RNeasy mini kit (Qiagen) per the manufacturer’s protocol sans further DNase I steps. Resulting RNA was quantified by NanoDrop (Thermo, ND-1000).

3.5.8 Microarray analysis of constitutive mutants.

Prior to microarray analysis, RNA was quantified using Qubit (Thermofisher) and assayed for quality on a Bioanalyzer using the RNA 6000 pico kit (Agilent). cDNA was prepared from total RNA using the Ovation Pico WTA System V2 (NuGEN). Samples were applied to custom GeneChips (Affymetrix) designed for P. aeruginosa PA14, B. thailandensis E264, Klebsiella pneumoniae, and Stenotrophomonas maltophilia genomes.

Runs were analyzed with Expression Console and Transcriptome Analysis Console v3.0 provided by Affymetrix.

3.5.9 cDNA synthesis and quantitative real-time PCR.

cDNA was generated using the SuperScript First-Strand Synthesis System for RT-

PCR (Invitrogen), following the manufacturer’s protocol for transcripts with high GC content. Transcripts were amplified using 1 µL NS5 primer and 2.5 µL of 100 ng/µL

RNA per sample.

Reactions for qRT-PCR consisted of 7.5 µL Q5 High-Fidelity 2x Master mix

(NEB), 1.6 µL 1x SYBR Green I (Thermofisher), 0.2 µL of each of the two 20 µM

116 primers used, 2.75 µL water, 0.75 µL DMSO, and 2 µL of cDNA. qPCR was performed using a BioRad QFX96 qPCR system. Samples were compared by calculating ΔΔCt values.

3.5.10 Growth of I0685 deletion and complemented strains in the presence of formaldehyde.

ΔI0685, ΔI0685 att::I0685 WT, and ΔI0685 att::I0685 N32G strains were constructed in a B. thailandensis ΔII1870 background to minimize the compensatory effects of other formaldehyde detoxification systems. Strains were grown overnight in

MOPS minimal media supplemented with 20 mM pyruvate and 5 mM glucose at 37oC.

Overnights were pelleted and washed 1x with 1 mL MOPS with 20 mM pyruvate, and resuspended in the same. Resuspended cells were diluted to an OD600 of 0.05 in either

MOPS plus 20 mM pyruvate or MOPS plus 20 mM pyruvate at 0.75 mM formaldehyde.

Cells were incubated at 37oC for 24 hours in a 24 well plate, 500 µL per well.

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3.6 Chapter 3 Figures

Figure 3.1 – GATR phylogeny based on full-length.

All full-length GATR amino acid sequences from B. thailandensis E264, B. pseudomallei, B. mallei, B. cenocepacia, PyrG (CTP synthetase) from E. coli, and FtrA orthologs from E. coli, C. crescentus, and Y. enterocolitica were aligned and a maximum likelihood, unrooted phylogenetic tree was constructed using 500 bootstrap replicates.

Bootstrap confidence values are displayed at branches.

Figure 3.2 – GATR phylogeny based on GATase1-like domain.

The amino acid sequences for bioinformatically predicted GATase1-like domains of the

GATRs of B. thailandensis E264, B. pseudomallei, B. mallei, B. cenocepacia, PyrG (CTP synthetase) from E. coli, and FtrA orthologs from E. coli, C. crescentus, and Y. enterocolitica were aligned and a maximum likelihood, unrooted phylogenetic tree was constructed using 500 bootstrap replicates. Bootstrap confidence values are displayed at branches.

Figure 3.3 – Growth of deletion strains on sole carbon sources.

The growth of WT and deletions strains on sole carbon sources at 37oC and was monitored by measuring OD600. The heat map represents averaged readings between the

24 and 48 hour time points. Darker purple indicates a higher optical density.

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Figure 3.4 – Growth of deletion strains on sole nitrogen sources.

The growth of WT and deletions strains on sole nitrogen sources at 37oC and was monitored by measuring OD600. The heat map represents averaged readings between the

24 and 48 hour time points. Darker purple indicates a higher optical density.

Figure 3.5 –II1850 (cdhR) regulates carnitine catabolism.

(A) The synteny of genes divergent from II1850 in B. thailandensis E264 and cdhR in P. aeruginosa PAO1 is compared. The divergent genes represent clear orthologs when sequences are compared and their arrangement is conserved between the two species. (B)

B. thailandensis E264 att::vector (WT) is able to grow on carnitine as a sole carbon source on MOPS minimal media plates. ΔII1850 att::vector is unable to utilize carnitine as a sole carbon source under the same conditions. The complementation of II1850 at the attTn7 site, under the control of its native promoter, restores the ability to grow on carnitine as a sole carbon source. This suggests that II1850 is an ortholog of CdhR and regulates the divergent genes in a similar manner.

Figure 3.6 – I1774 (argR) regulates arginine catabolism.

(A) Genes in the neighborhood of I1774 in B. thailandensis E264 are bioinformatically predicted to be involved in the catabolism and transport of arginine, similarly to those in the neighborhood of argR in P. aeruginosa PAO1. The synteny of genes is not perfectly

119 conserved between the species, but the similarity of predicted function is suggestive that

ArgR and I1774 are orthologous. (B) B. thailandensis E264 att::vector (WT) is able to grow on arginine as a sole carbon source on MOPS minimal media plates. ΔI1774 att::vector is unable to utilize arginine as a sole carbon source under the same conditions.

This defect may be complemented by placing the I1771-4 operon at the attTn7 site.

Figure 3.7 – Constitutive mutations in P. aeruginosa SouR overlaid on 3GRA.

The amino acid sequence of SouR from P. aeruginosa was overlaid on the crystal structure of its ortholog in P. putida, 3GRA. Constitutive mutants of SouR were sequenced and cognate amino acids in 3GRA were identified based on the overlay.

Highlighted mutations are as follows: P41S,T (red), D48G (orange), D76N (green), and

P185S (blue).

Figure 3.8 – Constitutive GbdR rescues growth on choline.

Similarly to the previously published ability of wild type gbdR to complement growth defects of ΔgbdR in P. aeruginosa on choline as a sole carbon source, ΔgbdR att::gbdR

E36G was able to rescue growth. This suggests that GbdR E36G is capable of folding properly and inducing GbdR controlled promoters.

Figure 3.9 – I0685 represses resistance to formaldehyde.

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ΔII1870 att::vector is capable of growing in the presence of 0.75 mM formaldehyde.

Deletion of ΔI0685 causes a slightly increased amount of growth, while complementation of the deletion with WT I0685 slightly reduces the amount of growth compared to wild type. Complementation of the deletion with I0685 N32G results in a striking decrease in the ability to tolerate formaldehyde compared to any of the other strains. Data is representative of three independent experiments.

Figure 3.10 – II1912 synteny in other Burkholderia.

Comparison of synteny for II1912 orthologs in multiple strains of B. thailandensis and one representative strain of B. pseudomallei. Schematic is approximate.

Figure 3.11 – FtrA is a potential regulator of cyanide detoxification.

(A) Alignment of amino acid sequences done as described for Figure 3.1, but nodes other than FtrA are collapsed for clarity. The FtrA node is highlighted in red. (B) Gene arrangement of II1760 and II1761. Synteny is identical for other Burkholderia species containing obvious II1760 orthologs. (C) Cyanide detoxification reaction catalyzed by rhodanese.

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3.7 Chapter 3 Tables

Table 3.1 – Strain list.

Table 3.2 – Plasmid list.

Table 3.3 – Primer list.

Table 3.4 – Burkholderia GATR homologs based on gene synteny.

This chart uses observed synteny and bioinformatic predictions of genes adjacent to

GATRs in B. thailandensis, B. pseudomallei, B. mallei, and B. cenocepacia in an attempt to predict potential orthologs. It also denotes if the divergent gene is predicted to be a DJ-

1 family member protein, a common feature of many putative GATRs.

Table 3.5 – Predicted constitutive mutants.

Predictions of residues that, when altered, may perturb the dimerization of the GATR and lead to constitutive activity.

Table 3.6 –GbdR E36G microarray results.

GbdR E36G induces expression of known GbdR regulon members in the absence of ligand. Cutoffs for data displayed are: fold effect ≥ 2 or ≤ -2, p value ≤ 0.05. Genes related to GB transport or catabolism are highlighted.

Table 3.7 – I0685 N32G microarray results.

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I0685 N32G downregulates expression of the glutathione independent formaldehyde dehydrogenase and its associated esterase. Cutoffs for data displayed are: fold effect ≥ 5 or ≤ -5, p value ≤ 0.05.

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CHAPTER 4:

DISCUSSION AND FUTURE DIRECTIONS

4.1 A brief summary of progress.

The first contribution of this work is the generation of an alternative model for the regulation of glycine betaine (GB) catabolism. In this new model, we identified multiple

AraC-family transcriptional regulators (ATRs) and characterized their phenotypes and target operons. The development of a model complementary to our original P. aeruginosa model provides the opportunity for to compare and contrast in order to gain new information about each. As part of this process, we identified and characterized three

ATRs in this pathway that are structurally related to the GATase1 enzymes, and represent a subfamily of ATRs referred to as Glutamine amidotransferase1 (GATase1)-containing

AraC-family transcriptional regulators (GATRs). The new regulatory model represents a scheme that better reflect the regulation found in Burkholderia, and may allow for assumptions of more distantly related bacteria that also possess these GATRs.

The second major contribution is the further characterization of GATRs as a family by presenting a methodology for perturbing the dimerization domain to yield constitutively active, ligand-independent mutants. This allows for the identification of target genes, whose potential functional predictions for which may inform us about the metabolic process that an unknown GATR is involved in and potentially ligands to which the GATR responds. Utilizing this method we identified I0685, a GATR that regulates

148 formaldehyde detoxification. The discovery of a GATR that functions as a repressor challenges our initial hypothesis that GATRs function strictly as activators, and opens the possibility that ligands for these regulators may not all be amine-containing compounds.

Simultaneously, these advancements have directly expanded our knowledge of the basic biology B. thailandensis. Several questions remain and our work has led to additional questions.

4.2 Remaining questions about the regulation of glycine betaine catabolism in B. thailandensis.

4.2.1 What are the possibilities for GbdR1/GbdR2 interaction?

We observed that GbdR1 and GbdR2 are both required for optimal induction of genes required for the transport and catabolism of GB. Furthermore, both GATRs can bind to the promoter region of the large GB catabolic operon (Figures 2.1, 2.5) (1). The presence of GbdR1 is an absolute requirement for expression of GbdR2 and GbdR2 increases the expression of GbdR1 (Table 2.1). This suggests that there is some manner of interaction between these two GATRs. However, the experiments described here were not sufficient to determine the particular nature of this relationship.

There are several possible explanations for the presence of two orthologs in this system. It is possible that there are unidentified half sites or overlapping binding sites for

GbdR2 within the portion of promoter used in our experiments. Such sites might allow

GbdR2 to regulate expression of GbdR1, as opposed to other ATRs which have been shown to autoregulate their own expression (2-4). Multiple binding sites might provide

149 an explanation for the banding pattern seen in electromobility shift assays incorporating

GbdR2 (Figure 2.5B).

We also observed that the N-terminal arm of GbdR2 is slightly longer than other

GATR N-terminal sequences. The presence of a longer N-terminal arm might offer an opportunity for regulation either through protein degradation or perhaps as a mechanical sensor for osmotic stress where its structure may be influenced by the physiological state of the cell (5, 6). Perhaps the arm affects the dimerization state by altering structural conformation as it does in the case of AraC (7). These possibilities might allow GbdR2 to act as a licensing factor for catabolism of GB under favorable conditions while preventing its breakdown during osmotic stress.

In order to determine the relationship between GbdR1 and GbdR2, there are several relatively simple approaches we could take. The first is to more carefully determine the binding site of each regulator. Our initial attempts to achieve this through

DNAase footprinting assays were unsuccessful, but there are many adjustments on the technique that could be applied. Alternatively, ChIP-seq might provide insights by identifying new target promoters, cumulatively from which binding sites and half sites which we were not able to detect might be derived (8).

The challenge in working with ATRs biochemically is thought to be due to a number of factors including their ability to oligomerize, potential folding issues when overexpressed protein is unbound, and potential non-specific binding to DNA that disrupts cell function during overexpression (9). However, much of this difficulty can be abrogated by expressing the domains individually, and focusing on just the C-terminal domain to determine the target binding site (10, 11). We could also repeat our RNA-seq

150 experiments under osmotically stressful conditions to discern if the apparent regulons of these regulators changes.

4.2.2 Likely influences on the regulation of GB catabolism in B. thailandensis that remain unaddressed.

Although the focus of our study has been on the GATRs involved in the regulation of GB catabolism, there are additional points of regulation in the pathway based on the model in P. aeruginosa that are likely to impact it. The breakdown of choline into GB is accomplished by the products of the betBA genes in P. aeruginosa, which are regulated by the TetR-family repressor, BetI (12, 13). These genes have also been identified in B. thailandensis, but it is not known if BetI impacts genes beyond betBA in this organism (14). Recently the expression of gbdR in P. aeruginosa was found to be under the direct control of BetI, as well as the global regulators NtrC and CrbB responsible for accessory nitrogen and carbon catabolism respectively (15). Notably,

CrbB has also been shown to be necessary for the expression of the operon containing the

GATR argR in P. aeruginosa (16). Nucleoid-associated proteins like H-NS, which are functionally similar to histones, are increasingly appreciated to play an important regulatory role in gene expression and have been shown to compete with ATRs for binding sites (17-19). Though not described, this is most certainly the case for B. thailandensis as well and it is possible that nucleoid-associated proteins impact the regulation of the GB catabolic pathway.

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Exploration of the additional regulators impacting this pathway could benefit from several approaches. First, the individual characterization of the binding sites of these regulators to enable the bioinformatic prediction of target promoters. Alternatively, deletion strains could be generated and used in combination with reporter plasmids similarly to our previous strategy to determine if they impact the expression of the GB catabolic operon (1). The findings could then be verified through ChIP to confirm physical interaction between the predicted binding sites and regulator. However, this approach might be hindered by the degree to which these global regulators can impact the cell and that may obfuscate the results.

4.2.3 How important is GB as an osmolyte and are there pools of related molecules in B. thailandensis?

Despite our focus on the importance of the GB catabolic pathway and our lab’s past interest in osmoprotection and molecular pools of osmolytes, we have not focused specifically on osmoprotection in B. thailandensis. The particular characteristics that predispose a given molecule to lend the cell resistance to osmotic stress, or for that matter barometric or thermal stress, are somewhat abstract. It is clear that such a compound must be a compatible solute, that is, the cell must be able to accumulate the molecule to high levels without disrupting the other biological processes and pathways necessary for life

(20). Ideally, the amount of these molecules would be readily controlled by means of transport or metabolic flux to compensate for a rapidly changing environment.

Typically for many bacteria, a sudden increase in hypertonic osmotic stress results

152 in the rapid uptake of K+ and production of glutamate, an often less than ideal countermeasure (21). The cell then accumulates and switches to the preferred osmolytes, while reducing the intracellular level of glutamate (22). Beyond GB and closely related compounds, other osmolytes found to be used by bacteria include trehalose, proline, ectoine, dimethylsulfoniopropionate, and others (23-28).

Previously our lab has utilized 13C nuclear magnetic resonance to examine the presence of molecular pools of choline and GB present during conditions of osmotic stress in different proteobacteria. As part of these experiments Burkholderia cepacia was observed to maintain a small choline pool and a larger GB pool, possibly functioning as a hedge against osmotic stress (29). B. thailandensis almost certainly maintains similar pools due to this being shown in B. cepacia and in addition to many other proteobacteria, but it has not been shown empirically. Preliminary experiments have also shown that B. thailandensis grown on choline is more tolerant to salt stress than when grown on pyruvate (unpublished). Presumably, these findings and the significant conservation of pathways resulting in the production of GB in Burkholderia indicate that examining GB induced osmoprotection in B. thailandensis would yield the same results. However, understanding the molecular mechanisms behind osmoprotection in B. thailandensis, and by extension its close relatives B. pseudomallei and B. mallei, will address pertinent biological questions. For instance, how does B. pseudomallei manage to endure environmental stress as the seasons changes from rainy to dry where it is endemic (30,

31)? Additionally, might GB related metabolism be important for interaction with plants

(33)?

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4.2.4 How closely is the transport and catabolism of choline-O-sulfate linked to

GbdR1 and GbdR2?

Choline-O-Sulfate (CS) catabolic operon is expressed in the presence of choline, dependent on the presence of both GbdR1 and GbdR2 (1). CS is used as a mechanism for sulfur storage in fungi, as well as an osmolyte by many plants (34, 35). Certain members of the Plumbaginaceae (a family of salt tolerant plants) preferentially utilize CS and β- alanine betaine as osmoprotectants over GB, likely due to the utility of using the generation of CS as a means to detoxify sulfur in the salt marsh environment (36). Many bacteria can utilize CS as an osmolyte or indirectly by converting it into GB, and in the case of B. thailandensis, subsequently using it as a carbon and nitrogen source (37, 38).

B. thailandensis possess genes for the transport and catabolism of CS located near the genes involved in the catabolism of GB. These genes encode a choline sulfatase (betC), a

CS substrate binding periplasmic transport protein (betD), a sulfate transporter (betE), and a divergently transcribed LysR-family transcriptional regulator (betR). This gene arrangement is broadly conserved in Burkholderia, particularly in the closely related B. pseudomallei and B. mallei. In Pseudomonas species, BetR has been shown to positively activate transcription of this operon in the presence of CS, but not choline or GB, and to negatively repress its own transcription in the absence of CS (39).

We have shown that B. thailandensis can utilize CS as either a sole carbon or nitrogen source (Figures 3.3 and 3.4). Studies from our own lab on the regulons of GbdR- like GATRs in P. aeruginosa have not suggested any such regulatory link exists between the CS catabolism and any of the GATRs (40, 41). It is conceivable that the upregulation of CS-related genes is due to read through transcription of the large GB catabolic operon,

154 which might continue to include the CS catabolic operon despite being interrupted by purU-2 and betR being transcribed in the opposite direction between the two (Figure

2.1C). Another possibility is that the CS catabolic operon or betR are regulated directly by either GbdR1 or GbdR2. The answer to this could be tackled by simple genetic techniques already utilized in our lab and would make an excellent project for an undergraduate.

The data obtained thus far suggest that B. thailandensis and other Burkholderia may have a more complex regulatory response to GB than initially appreciated, in a sense anticipating the environmental presence of CS through the presence of a downstream metabolite. The presence of pathways that utilize CS and related metabolites is logical in a bacterium that often resides in soil with an association to plants and fungi. The utilization of CS may provide an additional clue to the particular environmental niche where B. thailandensis and B. pseudomallei reside, given that its particular environmental reservoir is still unknown as discussed in Chapter 1.

4.2.5 The absence of SouR in B. mallei and potential effects on GB catabolism.

B. mallei lacks a sequence ortholog for SouR, the GATR that is responsible for the regulation of sarcosine catabolism, a downstream portion of the glycine betaine utilization pathway in B. thailandensis and P. aeruginosa (1, 40). An ortholog for the α- subunit of sarcosine oxidase is present in twenty seven out of forty two sequenced B. mallei strains while the γ-subunit ortholog can only be found in four of those strains (42).

Traces of the remaining sarcosine oxidase subunits and sdaA cannot be identified at all.

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Tellingly, putative transposases are uniquely located adjacent to the α-subunit suggesting that the sarcosine oxidase operon was truncated in the process of genetic reduction. This suggests that unlike B. pseudomallei and B. thailandensis, B. mallei is unable to utilize

GB or its catabolites as a carbon or nitrogen source due to a probable inability to degrade sarcosine.

Despite missing this piece of the pathway, B. mallei maintains orthologs for

GbdR1, GbdR2, and CdhR (Table 3.4, Figures 3.1 and 3.2). Also maintained are the operons that these GATRs regulate. B. mallei is considered to be derived directly from a particular isolate of B. pseudomallei that underwent significant genetic reduction due the proliferation of insertion sequence (IS) elements from transposons and phages (43-45).

This evolution and reduction continue through genomic recombination due to homology between IS elements (46). This genomic reduction likely led to B. mallei becoming an obligate parasite of mammalian cells, particular equines (47). It is possible that the transport systems for carnitine, choline, and GB retain importance for survival due to the role of GB as an osmoprotectant, but the acquisition of these compounds for usage as carbon and nitrogen sources is diminished due to the host environment being rich in other nutrient sources. It is possible that B. mallei possesses an alternative pathway for sarcosine degradation that would otherwise complete the larger pathway, but none are obvious.

Although B. mallei has never been isolated from the environment and is considered an obligate pathogen, it is possible to cultivate it under laboratory condition in minimal media (48). Thus it may be surprisingly simple to test if sarcosine may be utilized. Methods for genetic manipulation have been developed for B. mallei (49), and

156 many animal models for virulence have been published (50-54). It may be possible to use these methods to discern if carnitine or GB transport and catabolism play a role in fitness during B. mallei infection.

4.3 Deciphering the mechanism of GATRs

4.3.1 How do GATRs interact with RNA polymerase?

While most ATRs have been shown to function as activators, it remains unclear how GATRs that function as activators interact with RNA polymerase, if at all.

Activators may function through a number of mechanisms including class I and II activation, which involve direct interaction with RNA polymerase (55). In class I activation, the activator interacts with the α subunit C-terminal domain of RNA polymerase. In class II activation, the activator sits adjacent to the -35 site and interacts with the σ subunit. Additional mechanism include class III activation where multiple activators interact with different subunits of RNA polymerase and activators that change the promoter conformation such that RNA polymerase can recognize the -10 and -35 elements (56). Different ATRs have previously shown to act as class I or class II activators (57-60). Based on the position of the CdhR binding site relative to the caiX promoter in P. aeruginosa (61), it is most likely that GATRs are class I activators.

The type of activation may be ascertained in several different ways. Showing interaction by immunoprecipitation of RNA polymerase components would be informative. Improvements in our understanding of promoters targeted by GATRs may aid our understanding by determining the positioning of various GATRs with respect to

157 other elements of the promoter. Performing 5’ rapid amplification of cDNA ends (RACE) to determine the actual transcriptional start sites of GATR targeted promoters and determining GATR binding sites as previously mentioned would readily accomplish this.

Because we already have some evidence that I0685 functions as a repressor

(Figure 3.8, Table 3.7), it is possible that not all GATRs actually interact with RNA polymerase. However, it is still possible that I0685 N32G is simply locked in a repressive conformation. The location of binding sites within promoters often dictates whether a regulator functions as a repressor or an activator and can differ for the same regulator between promoters as with the ATR, MarA (62). However, it is possible that some findings will be transferable among more closely related GATRs, particularly those within the GbdR1 clade that appear to function specifically as activators (Figures 3.1 and

3.2).

4.3.2 A novel mechanism for ATR function.

The mechanism of GATRs, similar to other ATRs, likely revolves around a conformational change in the dimer to achieve either activation or de-repression. For

GATRs this conformational change is potentially derived from homology to GATases, which depend on dimerization for creation of an ammonia channel and subsequent conformation change to open the channel and transfer the ammonia to an associated synthase (63). Additional analysis of the 3GRA structure and the generation of new crystal structures in the presence and absence of ligand may provide insight into whether conformational changes from GATases are likely to be conserved in GATRs. We could

158 also incorporate our constitutive GATR mutants into these experiments to determine if the crystal structures, and subsequently the conformation of the predicted dimer, are altered.

In other ATRs such as AraC, changes in conformation arise in part from interaction of the N-terminal arm with bound ligand (7, 64-66). Due to the unconserved nature of the N-terminal arm in GATRs, it is unclear if it serves a similar function. N- terminally affinity tagged GATRs have been shown to bind DNA targets in vitro (1, 40,

41, 61), which might be expected to impact the functionality of the arm and subsequently the DNA binding properties of the protein. Future studies should employ strains with deletions of the N-terminal arm to help determine if the arms is a critical component of known GATRs. These apparent differences between GATR mechanism from previously described mechanisms of ATRs offers a new, additional model for ATR function.

4.3.3 The potential functions of unknown GATRs.

Several GATRs are divergently transcribed from putative DJ-1/Pfp1family proteins (Figure 1.7, II0513 and II1912), a family which includes GATase1 domain containing proteins. It is tempting to speculate if homology between the GATR and adjacent protein product might reflect commonalities between the ligand for the regulator and the substrate for the enzyme. One of these adjacent genes in B. thailandensis,

II0514, has recently been bioinformatically predicted to be involved in the degredation of caprolactam (42). Caprolactam is a cyclic amide used in the production of Nylon 6 and may be used as a carbon source by some bacteria (67).

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Other GATRs of unknown function are also divergently transcribed from their likely regulatory targets. II2197 is divergently transcribed from a predicted O- acetylserine sulfhydrylase, an enzyme that generates L-cysteine and acetate from bisulfide and O-acetyl-L-serine (68). Although our carbon and nitrogen screens included

O-acetyl-L-serine (Figure 3.3, 3.4), it is possible that there is no efficient transporter specific for O-acetly-L-serine and some bacteria are known to contain multiple O- acetylserine sulfhydrylases (69). II0282 is divergently transcribed from a gene encoding a predicted isochorismatase-family protein. Isochorismatase family proteins include a

GATase1 domain and catalyze the conversion of chorismate to 2,3-dihydroxybenzoate and pyruvate. Chorismate itself is an important precursor for aromatic amino acids, folate, vitamin K, salicylic acid, and siderophores (70, 71). A description of the relationship between II1760 and rhodanese is described in chapter 3 and between I1350 and capsule genes in Appendix A.

These potential functions greatly broaden the scope of metabolic processes which

GATRs are involved in. These insights also point to additional compounds to test the

GATR deletion strains against in future variations of our carbon- and nitrogen-source screens (Figures 3.3, 3.4).

4.4 Concluding remarks.

B. thailandensis has proven to be an excellent model for the exploration of

GATRs and we have identified the function of roughly half the GATRs in this organism.

Importantly, we have gained insight into a GATR outside of the GbdR clade (Figure 4.1).

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This has challenged some of our initial assumptions about this family of regulators, as we now understand they may act as either activators or repressors, and may not be limited to amine-containing compounds as ligands. With this new information, we can refine our methodology for generating constitutive mutants, gain understanding of the remaining

GATRs in B. thailandensis, and hopefully unravel the molecular mechanisms behind

GATR function.

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4.5 Chapter 4 Figures

Figure 4.1 – GATRs of known function in B. thailandensis.

This phylogenetic tree was constructed as described in chapter 3 of this thesis. GATRs with known functions are highlighted in purple. While most of the described GATRs are clustered together, notably I0685 is located on the far side of the tree. Thus, perhaps new insights into the remaining GATRs may be gleaned from the differences we find between the GbdR1-like clade and I0685.

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Appendix A - Biofilm formation in Burkholderia thailandensis.

A.1 Introduction.

Our previous investigation into the GATRs GbdR1, GbdR2, and SouR included an RNAseq, which included a tantalizing change in expression of yet another GATR and the adjacent operon (1). This GATR, I1350, exhibited changes in expression in the same range as the adjacent operon, which it may be a part of and we hypothesized that it might regulate. This particular operon is one of the major capsule operons in B. thailandensis, and the potential for a GATR to impact capsule expression and a possible connection with other GATRs piqued our curiosity.

B. pseudomallei possesses four major capsular polysaccharide gene clusters (CPS

I-IV), of which it shares three with B. thailandensis. Notably missing from B. thailandensis but present in B. mallei, is CPS-I. CPS-I is considered an important virulence determinant for both B. pseudomallei and B. mallei because it reduces the efficiency of opsonization in the blood stream, thus reducing phagocytosis (2-4).

Expression of CPS-III is increased in water, and may be associated with the ability of B. pseudomallei and B. mallei to persist in pure water for extended durations (5). CPS-II and

CPS-IV (which is adjacent to I1350 and the focus of our interest here) have no obvious effect on virulence and their roles are currently unknown (6). In addition to these, there are several smaller gene clusters relating to exopolysaccharide production, modification, and transport, some of which have been shown to correspond to biofilm production in B. pseudomallei (7). Thus far, current studies of the major CPS and potential link to biofilm

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formation are currently inconclusive since they may provide some redundant functionality (8).

The perturbation of genes that affect the outer-most layer of the cell and consequently biofilm formation was an attractive thread to pull. Like many other bacteria,

B. thailandensis and its relatives are known to form biofilms that contribute to resistance against drugs, desiccation, and predation (9). Therefore, we performed a series of experiments to examine potential morphological and biofilm phenotypes in our deletion strains.

A.2 Methods.

Construction of strains. The method of construction for these strains has been previously published (1), or described elsewhere in this thesis. Briefly, as per Thongdee et al. (10), primers were designed to amplify regions upstream and downstream of the region targeted for deletion. In tandem, a trimethoprim resistance marker enclosed by FLP recombinase sites was amplified with twenty base pair regions on the ends with homology to the amplified upstream and downstream fragments. Splice overlap extension

PCR was used to combine these fragments. Wild type B. thailandensis was grown overnight in DM media to induce natural competence, and were pelleted and combined with the splice overlap extension amplified product. Homologous recombination resulted in the generation of deletion strains that were selected on LB plates containing 100

µg/mL trimethoprim. The trimethoprim resistance marker was removed by transforming

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the trimethoprim resistant deletion strains with pFLPe2 by electroporation. pFLPe2 is a plasmid expressing FLP recombinase under the control of a rhamnose-inducible promoter as described in Choi et al. (11). After selecting transformants on LSLB plates with 0.2% rhamnose and 250 µg/mL zeocin incubated at 30oC, and subculturing them a second time on the same type of plate, they were struck for isolation on standard LB plates and grown at 42oC overnight in order to cure pFLPe2, which possesses a temperature sensitive origin. Colonies were then plated onto LB incubated at 37oC, LSLB with 250 µg/mL zeocin incubated at 30oC overnight to test for curing of pFLPe2, and LB with 100 µg/mL trimethoprim incubated at 37oC overnight to test for absence of the trimethoprim resistance marker. Trimethoprim sensitive deletion strains were confirmed by PCR and saved.

Spot plating and microscopy. Strains were struck out onto LB agar plates from -80oC stocks and incubated overnight at 37oC. For each strain, 3 mL LB broth was inoculated and incubated overnight at 37oC. Strains were diluted to an OD600 of 0.25 using LB broth, then 5 µL of each was spotted onto an R2B plate. Spots were allowed to dry for up to thirty minutes at room temperature before flipping the plate and incubating overnight at 37oC. Photographs of resulting macro-colonies were taken using a Leica MZ16 F stereomicroscope and Leica DC500 CCD camera at the University of Vermont

Microscopy Imaging Center.

Biofilm assay. The methodology for this assay was adapted from the popular microtiter

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biofilm assay popularized by George O’Toole’s lab at Dartmouth (12). Strains were struck out from -80oC stocks onto LB agar plates and incubated overnight at 37oC. For each strain, 3 mL LB broth aliquots were inoculated from the plates and incubated overnight at 37oC. At stationary phase, the overnights were diluted one to one hundred in

LB. Added 150 µL of diluted bacteria to five technical replicate wells each on a Costar

Serocluster 96 well “U” bottom, vinyl plate (REF 2797). Place plate in a small container with a moistened paper towel to act as a humidity chamber. Incubate at 37oC overnight.

Invert plate over waste container and shake gently. Rinse twice by gently submerging in distilled water. Tamp dry on paper towels. Add 180 µL of 0.1% crystal violet to each well and let sit for ten minutes. Invert plate over waste tray and shake gently. Rinse plate twice gently in distilled water, invert and tamp down on paper towels to dry. add 150 µL

80% ethanol to each well, wait five minutes for the crystal violet to solubilize, and read plate at OD550 on a Biotek Synergy 2.

RNAseq analysis. In previous work (1), B. thailandensis strains were examined to elucidate the putative regulons for GbdR1 and GbdR2. WT. ΔgbdR1, ΔgbdR2, and

ΔgbdR1ΔgbdR2 strains were grown in MOPS media with 20 mM pyruvate in the presence or absence of 1 mM choline. Inductions were done in duplicate and RNAseq was performed as previously described. Comparisons were filtered as follows: 1) WT + choline versus WT no choline log fold change (expressed as log2(X)) ≥ -2 or ≤ 2, WT + choline versus ΔgbdR2 + choline ≤-2 or ≥2 and significance ≤ 0.05, 3) WT + choline versus ΔgbdR1ΔgbdR2 + choline ≤-2 or ≥2 and significance ≤ 0.05. Applying these

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filters results in a comparison of genes that are up or down regulated in ΔgbdR2 and

ΔgbdR1ΔgbdR2 compared to the WT, but not due to the presence or absence of choline.

Presumably, the results include genes that might be responsible for the colony morphology and biofilm phenotypes. Data is available at NCBI’s Gene Expression

Omnibus, accession GSE81652.

A.3 Results and Discussion.

Our previous work investigating the regulons of GbdR1, GbdR2, and SouR, as well as their role in glycine betaine catabolism, hinted at some relationship between strains containing a deletion of gbdR2 and the uncharacterized GATR, I1350 (1).

However the nature of this putative relationship remained unclear and did not seem to directly relate the presence or absence of choline. In other instances, GATRs have been shown to impinge on the regulation of promoters primarily ascribed to another GATR, as in the case of GbdR and SouR in P. aeruginosa (13), as well as for GbdR and CdhR

(unpublished, Jamie Meadows). Thus, it was tempting to speculate that there might be some regulatory cross-talk between GbdR2 and I1350 in B. thailandensis. I1350 is a particularly interesting GATR because of synteny with a major capsule operon in B. thailandensis, possibly being transcribed as part of the same operon. The proximity of this GATR to CPS-IV (I1351 to I1362), as well as previous RNAseq data which showed changes in expression being relatively uniform across I1350 to I1362, make it attractive to consider that I1350 might be involved in the regulation of this capsule. This particular

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capsule operon is homologous to capsule IV in B. pseudomallei. Although capsule I of B. pseudomallei is primarily responsible for virulence, experiments in the Golden Syrian

Hamster model suggested that CPS-IV may have some contribution to virulence.

However, the role of CPS-IV with respect to the biology of B. pseudomallei or B. thailandensis is not well understood or particularly well studied due to being overshadowed by the clear importance of CPS-I.

In the course of working with B. thailandensis strains, it was noticed that some of the deletion mutants exhibited differential colony morphologies. This difference was particularly notable when a number of cells were spotted onto plates as opposed to colonies arising from single colony forming units. Spotting bacteria onto R2B plates consistently yielded a striking ‘stained glass’ morphology for ΔgbdR2 and

ΔgbdR1ΔgbdR2 strains, but not for WT or ΔgbdR1 strains (Figure 5.1 A). This morphological phenotype was also present in the ΔI1350. Unfortunately, none of the various GATR complementation strains we constructed altered this morphological phenotype (data not shown). This was true despite the GbdR1 and GbdR2 complementation strains having been shown to phenotypically complement defect in growth of gbdR1 and gbdR2 deletion strains when grown on choline or its derivative metabolites as sole carbon sources (1). The inability of clearly functional catabolic complementation to alter this morphological phenotype suggests that the hypothetical relationship between gbdR2 is incorrect, but the possibility existed that other explanations could account for the disparity.

Because of the potential of capsule modification to impinge upon biofilm

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formation due to alterations of the cell’s exterior surface, we opted to utilize the microtiter plate biofilm assay popularized by the O’Toole lab (12). Coincident with this project, many other GATR deletion strains had been produced (Table 5.1). We included the GATR deletion strains available to us because of the simplicity of the screen and achieved surprising results (Figure 5.1 B). Seven out of the eleven deletion strains appeared to have some level of defect in biofilm formation. Barring a massive level of cross-talk between these regulators, it became evident that the source of the biofilm defect was more likely due to other mutations, perhaps selected for by the process involved for subculturing and creating these strains.

Another possible explanation for the apparent biofilm defect in our various deletion strains was that the phenotype is due to a subpopulation within the wild type stock that contains the responsible mutation or that the unknown factor regulated by phase variation. In order to untangle these possible explanations, the wild type stock was struck on LB agar plates and isolated colonies were selected and subjected to our biofilm assay (Figure 5.2 A). This resulted in many clones exhibiting typical levels of biofilm formation, and several that exhibited a defect. Clones 9 and 10 were further subcultured and struck for isolation on LB agar plates. These subclones were again subjected to the biofilm assay and were found to yield phenotypes consistent with their parental clone

(Figure 5.2 B). Additional experimentation was consistent with these results (data not shown). The absence of any biofilm phenotype revertants in either direction makes it unlikely that phase variation is the cause of the phenotypes. Rather, the most likely explanation is that our original stock of wild type B. thailandensis E264 contains a

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subpopulation that exhibits a biofilm formation defect. This is critical to note as we intend to further investigate many of the GATR deletion strains we created for as of yet unknown phenotypes and were this not known, it could convolute our future results.

In order to derive some explanation for our observed phenotypes, we delved back into our old RNAseq data (Table 5.2). Because of the strains and conditions used, it was possible to eliminate genes expression changes that were specifically due to choline, which the gbdR1 and gbdR2 deletions impinge directly on. Also because both ΔgbdR2 and ΔgbdR1ΔgbdR2 expressed the colony morphology and biofilm deficient phenotypes, we could eliminate disparities between the two strains unrelated to these particular phenotypes and compare both to wild type. Doing so reveals two large operons that may be responsible for these phenotypes. The first, I1350-I1357, contains the capsule operon linked with the putative GATR regulating it, I1350, which initially captured our interest.

The second putative operon, I2674-I268, contains a putative two-component regulatory response system and genes that are bioinformatically predicted to be involved with the assembly of fimbriae. The altered expression of capsule and fimbriae genes is a logical source for the observed phenotypes, as they could both easily be imagined to affect cell to cell or cell to surface interactions. Both of these clusters are present in B. pseudomallei, although neither has been directly tied to virulence. There are other fimbriae genes that have been associated with both virulence in B. pseudomallei and biofilm formation in B. cenocepacia (14, 15). Our experiments with these strains utilizing the Madagascar hissing cockroach infection model did not suggest any impact on virulence (data not shown) (16). However, the question of why these genes are expressed differentially

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remains. Further investigation would require the construction of deletion and complementation strains to explain the phenotype at a minimum. At present, we are content with this explanation and subsequent genetic manipulation for use in our research in B. thailandensis E264 has been with a WT subclone confirmed for biofilm production.

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A.4 Appendix A Figures

Figure A.1 – Searching for a link between phenotype and genotype.

(A) Deletion strains of B. thailandensis E264 on R2B plates exhibited consistent morphological phenotypes. WT and ΔgbdR1 have topologically flatter, filled in centers.

ΔgbdR2, ΔgbdR1ΔgbdR2, and ΔI1350 exhibited an unusual ‘stained glass’ morphology.

(B) Available GATR deletion strains were tested for biofilm formation utilizing crystal violet staining. Surprisingly, seven out of the twelve strains tested exhibited a defect in biofilm formation.

Figure A.2 – Biofilm phenotype subculturing in B. thailandensis E264.

(A) Subclones of WT B. thailandensis E264 from our stock vile were isolated and subjected to crystal violet biofilm assay. Both biofilm positive and biofilm negative subclones were present. (B) Selected subclones nine and ten were struck out for isolation and secondarily subclones. These subclones were also subjected to crystal violet biofilm assaying, and their phenotypes were consistent with the subpopulation from which each was derived.

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A.5 Appendix A Tables

Table A.1 – Strain Utilized in Study.

A list of strains derived from Burkholderia thailandensis E264 utilized in the experiments described in this chapter. AN3, AN16, AN19, and AN126 were first described in previously published work (1).

Table A.2 – RNAseq Analysis Correlation.

Previously obtained RNAseq results were differentially filtered in order to seek a potential explanation for the phenotypes seen in ΔgbdR2 (AN19) and ΔgbdR1ΔgbdR2

(AN16) strains, but do not include changes in gene expression that are due to the presence of choline. Two sets of interesting genes were downregulated in relation to the

WT strain (MJ358). These included an operon involving capsule production and its presumptive GATR regulator, as well as an operon bioinformatically predicted to be involved in the assembly and transport of fimbriae. ‘m’ in between strain names signifies subtractive comparison. ‘.lfc’ signifies log fold change expressed as either positive or negative log2(x). ‘.p’ signifies the calculated significance of the correlation.

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A.6 References

1. Nock AM, Wargo MJ. 2016. Choline Catabolism in Burkholderia thailandensis Is Regulated by Multiple Glutamine Amidotransferase 1-Containing AraC Family Transcriptional Regulators. J Bacteriol 198:2503-2514. 2. DeShazer D, Waag DM, Fritz DL, Woods DE. 2001. Identification of a Burkholderia mallei polysaccharide gene cluster by subtractive hybridization and demonstration that the encoded capsule is an essential virulence determinant. Microb Pathog 30:253-269. 3. Reckseidler SL, DeShazer D, Sokol PA, Woods DE. 2001. Detection of bacterial virulence genes by subtractive hybridization: identification of capsular polysaccharide of Burkholderia pseudomallei as a major virulence determinant. Infect Immun 69:34-44. 4. Reckseidler-Zenteno SL, DeVinney R, Woods DE. 2005. The capsular polysaccharide of Burkholderia pseudomallei contributes to survival in serum by reducing complement factor C3b deposition. Infect Immun 73:1106-1115. 5. Reckseidler-Zenteno SL, Viteri DF, Moore R, Wong E, Tuanyok A, Woods DE. 2010. Characterization of the type III capsular polysaccharide produced by Burkholderia pseudomallei. J Med Microbiol 59:1403-1414. 6. Reckseidler-Zenteno SL, Moore R, Woods DE. 2009. Genetics and function of the capsules of Burkholderia pseudomallei and their potential as therapeutic targets. Mini Rev Med Chem 9:265-271. 7. Borlee GI, Plumley BA, Martin KH, Somprasong N, Mangalea MR, Islam MN, Burtnick MN, Brett PJ, Steinmetz I, AuCoin DP, Belisle JT, Crick DC, Schweizer HP, Borlee BR. 2017. Genome-scale analysis of the genes that contribute to Burkholderia pseudomallei biofilm formation identifies a crucial exopolysaccharide biosynthesis gene cluster. Plos Neglected Tropical Diseases 11. 8. Tseng BS, Majerczyk CD, Passos da Silva D, Chandler JR, Greenberg EP, Parsek MR. 2016. Quorum Sensing Influences Burkholderia thailandensis Biofilm Development and Matrix Production. J Bacteriol 198:2643-2650. 9. Fazli M, Almblad H, Rybtke ML, Givskov M, Eberl L, Tolker-Nielsen T. 2014. Regulation of biofilm formation in Pseudomonas and Burkholderia species. Environ Microbiol 16:1961-1981. 10. Thongdee M, Gallagher LA, Schell M, Dharakul T, Songsivilai S, Manoil C. 2008. Targeted mutagenesis of Burkholderia thailandensis and Burkholderia pseudomallei through natural transformation of PCR fragments. Appl Environ Microbiol 74:2985-2989. 11. Choi KH, Mima T, Casart Y, Rholl D, Kumar A, Beacham IR, Schweizer HP. 2008. Genetic tools for select-agent-compliant manipulation of Burkholderia pseudomallei. Appl Environ Microbiol 74:1064-1075. 12. O'Toole GA. 2011. Microtiter dish biofilm formation assay. J Vis Exp doi:10.3791/2437. 204

13. Willsey GG, Wargo MJ. 2015. Sarcosine catabolism in Pseudomonas aeruginosa is transcriptionally regulated by SouR. J Bacteriol doi:10.1128/JB.00739-15. 14. Allwood EM, Devenish RJ, Prescott M, Adler B, Boyce JD. 2011. Strategies for Intracellular Survival of Burkholderia pseudomallei. Front Microbiol 2:170. 15. Tomich M, Mohr CD. 2003. Adherence and autoaggregation phenotypes of a

Burkholderia cenocepacia cable pilus mutant. FEMS Microbiol Lett 228:287-297. 16. Fisher NA, Ribot WJ, Applefeld W, DeShazer D. 2012. The Madagascar hissing cockroach as a novel surrogate host for Burkholderia pseudomallei, B. mallei and B. thailandensis. BMC Microbiol 12:117.

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Appendix B – A prophage affecting B. thailandensis E264.

B.1 Introduction.

B. thailandensis strains, including E264, are known to harbor various phages that are capable of infecting B. mallei, B. pseudomallei, and other bacteria (1). The degree to which these phages have been categorized often is dependent on their immediate experimental utility. The status of B. mallei and B. pseudomallei as select agents has stimulated interest in the identification of phages which might potentially infect them, either as diagnostic tools or potentially as therapeutics due to their capacity for drug resistance (2). Additionally, the identification of phages capable of infecting B. pseudomallei in soil samples could be used as an indicator of the likely presence of B. pseudomallei, which is not always consistently isolated from soil where it is known to be present (3).

While generating B. thailandensis E264 ΔgbdR1ΔgbdR2ΔsouR during the course of exploration of glycine betaine catabolism, we noticed the emergence of a lytic phage in one of the two clones, AN206 (Figure B.1 A). Bacteria from the plate were re-struck unto two new LB plates, with one being struck using inoculum including presumptive plaques and the other struck using inoculum seemingly clear of any plaques. The plate struck using inoculum was clearly riddled with phage (left) while the other plate contained no evidence of plaques (right) (Figure B.1 B). The initial preparation of phage was collected from this left plate by adding 10 mL of 10 mM Tris pH = 7.4 to the plate to rinse its surface, then filter sterilizing the collected liquid with a 0.22 µ filter. Adding 5

µL drops to an LB plate inoculated with a fresh lawn of wild type B. thailandensis E264

206 resulted in a large, totally cleared zone after an overnight incubation at 37oC (Figure B.1

C). These findings suggested the presence of a phage, which we have designated as

“There are some who call me Tim” or φTim for short (4). Because of the interest in phages affecting Burkholderiales, we endeavored to identify the phage.

B.2 Methods.

Amplification of bacteriophage. B. thailandensis E264 was grown overnight at 37oC in 3 mL LB media. It was then diluted by adding 1 mL of overnight to 50 mL LB, add 10 µL of phage, and incubated overnight at 37oC. Cells and debris were pelleted by spinning at

2500 rpm for 15 minutes in a 4.2 rotor. The supernatant was filter sterilized using a .22

µm filter.

Rough bacteriophage DNA isolation. To 5 mL of filtered bacteriophage, add 10 µL

DNase I (NEB) and incubate on ice for 1 hour to eliminate the majority of carry-over bacterial DNA. The filtrate was aliquoted to ten 1.5 mL microcentrifuge tubes, 300 µL each. To each tube, 0.5 µL glycogen, 33 µL 3 M NaOAc pH = 5.2, and 300 µL phenol chloroform isoamyl alcohol (50:49:1) were added. The tubes were vortexed thoroughly, then centrifuged for four minutes at 14,000 rpm in a table top microcentrifuge at room temperature. The aqueous phase was saved to a clean 1.5 mL microcentrifuge tube, to which 900 µL ethanol was added. Tubes were mixed by inversion and stored at -80oC overnight. Thawed samples were then spun at 14,000 rpm for 30 minutes at 4oC. The pellets were washed with 800 µL 80% ethanol, then air dried for five minutes. Each pellet was resuspended in 25 µL of water and pooled.

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Cloning of phage DNA into pGEM-T-Easy vector. 10 µL of phage DNA was combined with 5 µL cutsmart buffer (New England Biolabs (NEB)), 33 µL water, and 2 µL AluI

(NEB #R0137S) for 40 minutes at 37oC. The digestion was stopped by incubating at

80oC for 5 minutes. 0.5 µL Taq polymerase and 1 uL 10 mM dATP were added and the reaction, and it was incubated for 20 minutes at 72oC. The reaction was cleaned up using a GeneJET Gel Extraction and DNA cleanup Micro Kit following the standard protocol for reaction cleanup (Thermo Scientific #K0832). The AluI cut, adenylated phage DNA was then ligated into pGEM-T-Easy (Promega #A1360) following the manufacturers protocol. The ligation reaction was used to transform chemically competent Escherichia coli DH5α cells and selected on LB plates with 100 µg/mL carbenicillin. Plasmid DNA from clones was isolated using a GeneJET Plasmid Miniprep Kit (Thermo Scientific

#K0503). Plasmid DNA was screened for inserts by digesting with EcoRI-HF (NEB

#R3101), and five clones positive for inserts were sent for sequencing using the M13F primer.

B.3 Results and Discussion.

Sequencing inserts derived from a crude preparation of phage DNA resulted in nBLAST hits exactly matching sections of II1331, II1332, and II1364 found in B. thailandensis E264 (Table B.1), indicating that we have isolated the lytic form of a prophage. These genes are part of a previously recognized prophage-like genomic island that was not observed to be active in E264, but would be categorized as a member of the

Myoviridae (5). The Myoviridae are a group of non-enveloped bacteriophages with

208 contractile tails, the most common example being the famous bacteriophage T4 (6). This particular genomic island was present in roughly 25% of a survey of 29 strains of environmentally isolated B. thailandensis (7). BLAST results for the inserts homologous to II1331 and II1332 suggested close similarity to phages in B. pseudomallei ((φE122, accession CP000624), (φX216, accession JX681814), (Burkholderia phage BEK, accession CP008753), (φ52237, accession DQ087285)) and in B. thailandensis E202

(φE202, accession CP000623), and B. cenocepacia C6433 (Burkholderia phage KS14, accession HM461982 (8)).. The size of this phage genome is predicted to be between 35 and 38 kilobases depending on the bioinformatics parameters, and in our hands the prepared genomic DNA appeared as a single prominent band in this approximate size range (data not shown).

Interestingly, these similar phages do not contain sequences homologous to our

II1364 insert, although this gene falls within the confines of the bioinformatically predicted prophage-like island (5). II1364 is predicted to code for a DNA methyltransferase, which are often used to protect phage DNA from cleavage by host endonucleases (9). Although it is possible this phage may have been previously incidentally isolated and tested for its ability to infect B. pseudomallei or B. mallei without being further analyzed and annotated, it does not appear that this phage has been previously sequenced and submitted to NCBI apart from being a part of the B. thailandensis E264 genome. Further characterization of this phage may reveal it to be another useful tool in the study of Burkholderiales.

B.4 Appendix B Figure

209

Figure B.1 – A bacteriophage infecting B. thailandensis E264. (A) Plaques were initially observed when plating a particular clone of a deletion strain, the plate on the left side of the panel. (B) The plate with apparent plaques was re-struck onto two new LB plates, one with inoculum including the plaque (left) and one with inoculum seemingly free of plaques (right). (C) An LB plate was struck for confluency with B. thailandensis E264, and a 5 µL drop of filter sterilized phage preparation was added with a 5 µL drop of filtered 10 mM Tris pH = 7.4 as a negative control. The plate was then incubated overnight at 37oC.

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B.5 Appendix B Table

Table B.1 – Sequencing results for cloned inserts. Sequencing results were subjected to nBLAST and bioinformatic predictions were added to the table. Highlighted entries indicate genes that may be phage-associated.

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B.6 References

1. Woods DE, Jeddeloh JA, Fritz DL, DeShazer D. 2002. Burkholderia thailandensis E125 harbors a temperate bacteriophage specific for Burkholderia mallei. J Bacteriol 184:4003-4017. 2. Guang-Han O, Leang-Chung C, Vellasamy KM, Mariappan V, Li-Yen C, Vadivelu J. 2016. Experimental Phage Therapy for Burkholderia pseudomallei Infection. PLoS One 11:e0158213. 3. Withatanung P, Chantratita N, Muangsombut V, Saiprom N, Lertmemongkolchai G, Klumpp J, Clokie MR, Galyov EE, Korbsrisate S. 2016. Analyses of the Distribution Patterns of Burkholderia pseudomallei and Associated Phages in Soil Samples in Thailand Suggest That Phage Presence Reduces the Frequency of Bacterial Isolation. PLoS Negl Trop Dis 10:e0005005. 4. Coca-Cola Company Collection (Library of Congress), LC Purchase Collection (Library of Congress). 1974. Monty Python and the holy grail. Almi Cinema 5, Columbia Pictures Home Entertainment, S.l. United States.

5. Ronning CM, Losada L, Brinkac L, Inman J, Ulrich RL, Schell M, Nierman WC, Deshazer D. 2010. Genetic and phenotypic diversity in Burkholderia: contributions by prophage and phage-like elements. BMC Microbiol 10:202. 6. Karam JD, Miller ES. 2010. Bacteriophage T4 and its relatives. Virol J 7:293. 7. Yu YT, Kim HS, Chua HH, Lin CH, Sim SH, Lin DX, Derr A, Engels R, DeShazer D, Birren B, Nierman WC, Tan P. 2006. Genomic patterns of pathogen evolution revealed by comparison of Burkholderia pseudomallei, the causative agent of melioidosis, to avirulent Burkholderia thailandensis. Bmc Microbiology 6. 8. Lynch KH, Stothard P, Dennis JJ. 2010. Genomic analysis and relatedness of P2-like phages of the Burkholderia cepacia complex. BMC Genomics 11:599. 9. Murphy J, Mahony J, Ainsworth S, Nauta A, van Sinderen D. 2013. Bacteriophage orphan DNA methyltransferases: insights from their bacterial origin, function, and occurrence. Appl Environ Microbiol 79:7547-7555.

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